Download Signaling and Communication in Plant Symbiosis

Signaling and Communication in Plants
Silvia Perotto
František Baluška Editors
Signaling and
Communication in
Plant Symbiosis
Signaling and Communication in Plants
Series Editors
František Baluška
Department of Plant Cell Biology, IZMB, University of Bonn, Kirschallee 1, D-53115
Bonn, Germany
Jorge Vivanco
Center for Rhizosphere Biology, Colorado State University, 217 Shepardson Building,
Fort Collins, CO 80523-1173, USA
For further volumes:
http://www.springer.com/series/8094
.
Silvia Perotto • František Baluška
Editors
Signaling and Communication
in Plant Symbiosis
Editors
Dr. Silvia Perotto
Università di Torino
Dipto. Biologia Vegetale
Viale P. A. Mattioli 25
10125 Torino
Italy
[email protected]
Dr. František Baluška
Universität Bonn
Inst. Zelluläre und Molekulare
Botanik (IZMB)
Kirschallee 1
53115 Bonn
Germany
[email protected]
ISSN 1867-9048
e-ISSN 1867-9056
ISBN 978-3-642-20965-9
e-ISBN 978-3-642-20966-6
DOI 10.1007/978-3-642-20966-6
Springer Heidelberg Dordrecht London New York
Library of Congress Control Number: 2011938275
# Springer-Verlag Berlin Heidelberg 2012
This work is subject to copyright. All rights are reserved, whether the whole or part of the material is
concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting,
reproduction on microfilm or in any other way, and storage in data banks. Duplication of this publication
or parts thereof is permitted only under the provisions of the German Copyright Law of September 9,
1965, in its current version, and permission for use must always be obtained from Springer. Violations
are liable to prosecution under the German Copyright Law.
The use of general descriptive names, registered names, trademarks, etc. in this publication does not
imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use.
Printed on acid-free paper
Springer is part of Springer Science+Business Media (www.springer.com)
Preface
Because of their photosynthesis, plants are the major primary producers in terrestrial ecosystems and a precious source of organic carbon for all their microbial
symbionts. Biotrophic microbial symbionst derive nutrients from the tissues of the
living host, and they often colonise the plant cells where they form intracellular
structures that are the site of nutrient uptake and exchange, as well as of
interorganismal signaling and communication. To colonise the plant tissues via
balanced endosymbiosis, all biotrophic microbes, irrespective of their trophic
strategy, need to overcome the plant defense responses through an exchange of
molecular signals.
Plants are unique as they are able to associate with both prokaryotic and
eukaryotic microbes and establish with them well-balanced symbiotic interactions
that range from mutualism to antagonism. No other multicellular organisms give
rise to this variety of symbiotic interactions. Plants must be able to discriminate
between mutualistic micro-organisms that may exchange organic carbon for essential nutrients such as nitrogen and phosphorus, thus promoting plant growth, and
antagonistic pathogens that are only detrimental and cause disease. As this volume
is making clear, a fine tuning of the signals in plant symbioses is very complex and
still only partially understood.
This volume provides overviews of the current knowledge on a variety of
symbiotic systems. The first section is dedicated to signalling during the formation
of mutualistic symbioses in legume plants. Legume plants have been pivotal to
understand the genetic bases of symbioses and the comparison between nodule and
arbuscular mycorrhizal (AM) symbioses has revealed a common symbiosis (SYM)
signalling pathway leading to intracellular accommodation of fungal and nitrogen
fixing bacterial endosymbionts. The recent discovery that AM fungi secrete symbiotic signals that resemble rhizobial lipochito-oligosaccharides (Maillet et al. 2011)
brings the similarities between these two symbioses even further. Despite the
importance of legumes as model systems to study mutualistic symbioses, there is
a great diversity of plant-microbe interactions that involve nitrogen fixing bacteria
others than rhizobia, and a variety of other mycorrhizal and endophytic fungi. Some
of the chapters in this book provide current knowledge on these diverse
v
vi
Preface
interactions, and witness the progress in the unravelling of genetic determinants in
plant-microbe signalling, and the impressive amount of data emerged from the use
of both genomic and post-genomic approaches. The last section of the book is
focused on the interaction of plants with antagonistic biotrophs, ranging from
filamentous fungi to oomycetes, and to nematodes.
Torino
Bonn
Silvia Perotto
Frantisek Baluska
Reference
Maillet et al. 2011. Fungal lipochitooligosaccharides symbiotic signals in arbuscular mycorrhiza.
Nature, 469:58–63
Contents
The Role of Diffusible Signals in the Establishment of Rhizobial
and Mycorrhizal Symbioses . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
J. Benjamin Miller and Giles E.D. Oldroyd
Infection of Lotus japonicus Roots by Mesorhizobium loti . . . . . . . . . . . .
Katharina Markmann, Simona Radutoiu, and Jens Stougaard
Signalling and the Re-structuring of Plant Cell Architecture
in AM Symbiosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Andrea Genre
1
31
51
Signalling and Communication in the Actinorhizal Symbiosis . . . . . . . .
Claudine Franche and Didier Bogusz
73
Signalling in Cyanobacteria–Plant Symbioses . . . . . . . . . . . . . . . . . . . .
David G. Adams and Paula S. Duggan
93
Signalling in Ectomycorrhizal Symbiosis . . . . . . . . . . . . . . . . . . . . . . . . 123
Judith Felten, Francis Martin, and Valérie Legué
Signalling in the Epichloe¨ festucae: Perennial Ryegrass Mutualistic
Symbiotic Interaction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 143
Carla Eaton, Milena Mitic, and Barry Scott
Plant Infection by Biotrophic Fungal and Oomycete Pathogens . . . . . . . 183
Pamela H.P. Gan, Peter N. Dodds, and Adrienne R. Hardham
Compatibility in Biotrophic Plant–Fungal Interactions: Ustilago
maydis and Friends . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 213
Kerstin Schipper and Gunther Doehlemann
Compatible Plant-Root Knot Nematode Interaction and Parallels
with Symbiosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 239
Bruno Favery, Michaël Quentin, and Pierre Abad
Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 259
vii
The Role of Diffusible Signals
in the Establishment of Rhizobial
and Mycorrhizal Symbioses
J. Benjamin Miller and Giles E.D. Oldroyd
Abstract The roots of at least 80% of all angiosperms are able to engage in
symbiotic relationships with arbuscular mycorrhizal (AM) fungi of the group
Glomeromycota in order to derive macro- and micro-nutrients from the environment (Brachmann and Parniske, PLoS Biol 4:e239, 2006). Legume roots also form
a unique symbiosis with rhizobia in order to derive fixed nitrogen. The establishment of both of these symbioses depends upon signalling between the plant host and
the microorganism, of which a number of diffusible signals are essential. Here we
discuss the synthesis and role of these diffusible signals for the establishment of
both rhizobial and mycorrhizal symbioses.
1 Introduction: A Molecular Dialogue Between Host Plant
and Symbiont
In order for symbioses to be established between host and symbiont it is necessary
that tightly regulated communication occurs. In the case of symbiotic interactions
between plant roots and microorganisms in the rhizosphere, the plant must attract
and promote the symbiotic partner to interact with its root, whilst in turn the
microorganism must respond to distinguish itself as symbiotic rather than pathogenic and subsequently gain regulated entry into the root. This results in a situation
where each organism is required to participate in an elaborate communication in
order to allow the establishment and progression of symbiosis. The term “molecular
dialogue” was originally coined to describe this communication which occurs
between the roots of legumes and rhizobia (Denarie et al. 1993), and is also suitable
when considering the interaction between mycorrhiza and host plants.
J.B. Miller • G.E.D. Oldroyd (*)
Department of Disease and Stress Biology, John Innes Centre, Norwich Research Park,
Norwich, NR4 7UH, UK
e-mail: [email protected]
S. Perotto and F. Baluska (eds.), Signaling and Communication in Plant Symbiosis,
Signaling and Communication in Plants 11, DOI 10.1007/978-3-642-20966-6_1,
# Springer-Verlag Berlin Heidelberg 2012
1
2
J.B. Miller and G.E.D. Oldroyd
2 Diffusible Signals During Nodulation
During the establishment of the legume-Rhizobium symbiosis, plant roots release
flavonoids which are perceived by rhizobial bacteria. This perception leads to the
induction of rhizobial nod genes which encode proteins required for the synthesis of
Nodulation (Nod) factors. Nod factors are secreted by rhizobia and upon their
detection act as signalling molecules to the host plant (Oldroyd and Downie
2006, 2008). Nod factors are essential for nodulation and are important mediators
of host-range specificity.
2.1
Flavonoids
The earliest component of the molecular dialogue between legumes and rhizobia is
the synthesis of flavonoids by the plant root. These are constitutively produced
polyaromatic compounds that are perceived by rhizobia via the NodD protein (Peck
et al. 2006). This LysR-type transcriptional regulator then promotes Nod factor
biosynthesis by activating the transcription of nod genes (Fisher and Long 1993).
2.1.1
Structure and Synthesis
Flavonoids are diverse polyaromatic secondary metabolites consisting of a 15-carbon skeleton and are formed from a branch of the phenylpropanoid pathway. The
first committed step of flavonoid biosynthesis is catalysed by chalcone synthase;
this reaction involves the condensation of 4-coumaroyl-CoA with three molecules
of malonyl-CoA to form a chalcone flavonoid precursor (Fig. 1). This precursor
feeds into further biosynthetic reactions which yield either 5-deoxyflavonoids or
5-hydroxyflavonoids. Flavonoids can be further sub-classed into flavonoids and
isoflavonoids according to whether the phenyl group is attached to C2 or C3
(as with the flavonol kaempferol or the isoflavone daidzein, respectively; Fig. 1).
Flavonoid production is ubiquitous in plants and these compounds are typically
associated with plant defence responses, in addition to lignin and anthocyanin
production (Winkel-Shirley 2001). The first proof for the role of root-exuded
flavonoids during symbiosis was the induced expression of Sinorhizobium meliloti
nodulation genes (nodABC) by luteolin, a flavone (Peters et al. 1986), and the
induction of Rhizobium trifolii nod genes by flavones from clover (Redmond et al.
1986). Subsequent research has identified several other flavonoids involved in the
nodulation signalling of many plant species (reviewed by Broughton et al. 2000).
It has been noted that some flavonoid structures are only produced by particular
plants; for example, isoflavonoid production is limited to the Papilionoideae
(or Faboideae) subfamily of the Leguminosae (Dixon et al. 2002). This diversity
of flavonoid production has been associated with determining, at least in part, the
The Role of Diffusible Signals in the Establishment of Rhizobial and Mycorrhizal
3
L-phenylalanine
Phenylalanine ammonia lyase
trans-cinnamic acid
Cinnamic acid 4-hydroxylase
p-coumaric acid
4-coumarate:CoA ligase
4-coumaroyl-CoA
LIGNINS
Chalcone synthase
Chalcone reductase
Naringenin chalcone
Chalcone isomerase (type I + II)
Isoliquiritigenin
Chalcone isomerase (type II)
5-deoxyflavonoids
5-hydroxyflavonoids
Liquiritigenin
Naringenin
(Flavanone)
(Flavanone)
Isoflavone synthase
Flavanone 3-hydroxylase
Flavone synthase II
Dihydroxyflavone
Daidzein
(Isoflavone)
Genistein
(Isoflavone)
OH
Daidzein
HO
Dihydrokaempferol
(Dihydroflavonol)
Flavonol synthase
Kaempferol (Flavonol)
O
HO
2
O
2
3
3
OH
O
OH
OH
O
Kaempferol
ANTHOCYANINS
CONDENSED TANNINS
Fig. 1 Biosynthesis of flavonoids. Partial diagram of the phenylpropanoid pathway. Enzymes are
depicted in italics; 5-deoxyflavonoids and 5-hydroxyflavonoids are denoted by shaded boxes;
compounds in dashed boxes represent major side branches of the pathway. Inset shows chemical
structures of two example flavonoids isolated from root exudates. Figure adapted from Shaw et al.
(2006), Subramanian et al. (2007), Winkel-Shirley (2001) and Zhang et al. (2009)
specificity of Rhizobium responses (Cooper 2007; Gibson et al. 2008); for instance,
some flavonoids will act as nod gene inducers in one context, but as repressors in
another (Firmin et al. 1986).
2.1.2
Biological Activity
Two distinct roles for flavonoids and their related molecules have been suggested in
the context of early communication during the legume-Rhizobium symbiosis: (1)
they are involved in chemotaxis responses of rhizobia; (2) they modulate nod gene
expression of rhizobia (Shaw et al. 2006).
It is essential that rhizobia present in the rhizosphere are attracted to host roots in
order for the symbiosis to be formed. Plant roots secrete many compounds into the
rhizosphere and in doing so modulate microorganism populations (Dennis et al.
2010). Indeed, it has been estimated that 27% of carbon allocated to roots is
4
J.B. Miller and G.E.D. Oldroyd
deposited in the rhizosphere (Jones et al. 2009). Legumes must therefore selectively
encourage the chemotaxis of rhizobia and flavonoid secretion is important in this
process. Positive chemotaxis of S. meliloti was specifically demonstrated with
the flavone luteolin which is produced by the S. meliloti host Medicago sativa;
this response did not occur with naringenin or apigenin, two closely related
flavonoids not produced by M. sativa (Caetano-Anolles et al. 1988). Additionally,
flavonoid production in host roots is increased upon the addition of compatible
Rhizobium species (Schmidt et al. 1994). This increased flavonoid synthesis was
dependent upon Nod factor structure, implying a positive feedback loop in the
regulation of flavonoid production specifically in legume–rhizobia interactions
which are symbiotically favourable.
Arguably the most important and most well-characterised rhizobial response to
flavonoids is the induction of nod gene expression. At least 30 nod gene inducing
flavonoids have been identified, and their activity is typically in the low micromolar
to nanomolar range (Cooper 2004). The exact mechanism by which flavonoids are
perceived in rhizobia is unclear; however, the importance of NodD proteins is
apparent. NodDs are transcriptional regulators of the LysR-type family and primarily control the expression of nod genes, which are responsible for Nod factor
biosynthesis (Sect. 2.2). Rhizobia usually contain one to five NodD homologues,
depending on species; for example, S. meliloti contains three NodDs which share
greater than 77% amino acid identity (Honma and Ausubel 1987; Peck et al. 2006).
NodDs bind to conserved 55 bp DNA sequences of the promoter region of inducible
nod genes, the so-called nod box, and in doing so induce a bend in the DNA (Fisher
and Long 1993). This DNA bending appears to sharpen upon appropriate flavonoid
treatment, resulting in subsequent RNA polymerase binding and thereby activating
transcription (Chen et al. 2005).
Much genetic evidence suggests that NodD is involved in flavonoid perception.
NodD is necessary and sufficient for nodC expression in the presence of flavonoids
(Mulligan and Long 1985). NodD from Rhizobium leguminosarum bv. viciae
localises to the cytoplasmic membrane (Schlaman et al. 1989), which is also
where the flavonoid inducer naringenin accumulates (Recourt et al. 1989). Point
mutation of NodD proteins extends nod gene expression to include flavonoids
which are usually non-inducing (Burn et al. 1987; McIver et al. 1989). Together
with additional research this has led to the suggestion that NodD controls rhizobial
responses to flavonoids in a species-specific fashion (Horvath et al. 1987; Spaink
et al. 1987; Zaat et al. 1989). However, the direct biochemical interaction of NodD
and flavonoids has been difficult to prove, although more recent work has begun to
demonstrate this interaction (Li et al. 2008; Peck et al. 2006). The work of Peck
et al. (2006) has importantly shown that inducing and non-inducing flavonoids
promote binding of S. meliloti NodD1 to the nod box, suggesting that competitive
inhibition between inducing and non-inducing flavonoids may be important in
regulating nod gene expression and thus nodulation efficiency. In vivo binding of
S. meliloti NodD1 to the nod box upon luteolin treatment also requires the activity
of the chaperonin GroEL (Ogawa and Long 1995; Yeh et al. 2002).
The Role of Diffusible Signals in the Establishment of Rhizobial and Mycorrhizal
5
Flavonoid induction of NodD via the nod box has been well characterised, and
14 of the 16 genes required for Nod factor biosynthesis are regulated in this manner
in Sinorhizobium sp. strain NGR234 (Freiberg et al. 1997; Kobayashi et al. 2004).
The promoters of many other Rhizobium genes also contain nod boxes, as
demonstrated in NGR234 which responds by increasing transcription of 147 open
reading frames upon daidzein treatment (Perret et al. 1999). Indeed, flavonoid
treatment can also act to repress gene expression (Firmin et al. 1986), for example
coumestrol and medicarpin, flavonoids secreted by M. sativa roots, repress nodC
expression in S. meliloti (Zuanazzi et al. 1998). Different flavonoids therefore play
different roles to positively and negatively regulate nod gene expression in Rhizobium species.
Transient increases in intracellular calcium in R. leguminosarum bv. viciae have
recently been detected upon treatment with flavonoid inducers (Moscatiello et al.
2010). These calcium transients were NodD independent, suggesting that an
additional flavonoid-perception mechanism remains to be characterised in
R. leguminosarum bv. viciae. Flavonoid non-inducers did not activate the calcium
response in R. leguminosarum bv. viciae (Moscatiello et al. 2010), therefore
this alternative flavonoid-perception mechanism must be specifically activated
by only flavonoid inducers. It will therefore be interesting to know the exact
interplay between different flavonoids, NodD and calcium in the role of nod gene
induction.
Silencing of enzymes involved in the biosynthesis of flavonoids has confirmed
the importance of these secondary metabolites in establishing symbioses with
rhizobia. Medicago truncatula plants silenced for chalcone synthase, chalcone
reductase and flavone synthase II expression show decreased or no nodulation
with S. meliloti (Wasson et al. 2006; Zhang et al. 2009). Similar silencing
experiments in Glycine max also show decreased nodulation (Subramanian et al.
2006). These results are consistent with the importance of flavonoids in activating
Nod factor biosynthesis, although it has also been suggested that auxin transport in
host roots may be regulated by flavonoids and this has been proposed to be involved
in nodule organogenesis (Subramanian et al. 2007). However, the relative importance of flavonoids activating Nod factor biosynthesis versus regulating auxin for
nodule organogenesis remains to be resolved.
Non-flavonoid diffusible signals are also produced by legumes and perceived by
rhizobia, although their role appears to be relatively minor due to the higher
concentrations required for biological activity (Brencic and Winans 2005; Cooper
2007). The first non-flavonoid compounds to be identified were the betaines
trigonelline and stachydrine from M. sativa which activated nod gene expression
in S. meliloti (Phillips et al. 1992). Jasmonates also stimulate nod gene expression in
R. leguminosarum (Rosas et al. 1998) and Bradyrhizobium japonicum (Mabood
et al. 2006), whilst B. japonicum nod gene induction has also been described with
xanthones (Yuen et al. 1995). Interestingly, simple phenolics from wheat, such as
vanillin and isovanillin, are also able to act as nod gene inducers in Sinorhizobium
sp. strain NGR234 (Le Strange et al. 1990).
6
J.B. Miller and G.E.D. Oldroyd
2.2
Nod Factors
Upon perception of flavonoids, rhizobial NodD proteins induce nod gene transcription. Some of these nod gene products are enzymes involved in the production of a
suite of lipochitooligosaccharides (LCOs) called Nod factors. The first Nod factor
structure to be presented was that of S. meliloti (Fig. 2a; Lerouge et al. 1990). Nod
factors have a generalised structure consisting of a chitin backbone of usually three
to five b-1,4-linked N-acetlyglucosamine residues to which additional decorations
and substituents are added, including an acyl (fatty acid) chain at the non-reducing
terminus. These decorations vary between Rhizobium strains and may include the
addition of acetyl, methyl, sulfate and sugar moieties (Fig. 2b). A single species of
rhizobia may produce several different Nod factors; for example, Rhizobium tropici
CIAT899 produces 52 different LCOs at acidic pH and 29 LCOs at neutral pH, yet
only 15 structures are common to both growth conditions (Morón et al. 2005).
a
O
NodL
NodH, NodP, NodQ
H 3C
OH
O
O
HO
HO
NH
O
O
HO
O
O
b
non-reducing
terminus
O
N
Acyl
H3C
O
n
NodA, NodB, NodC
O
R1
O R4
OH
OH
O R3
R2
R2
OH
NH
H3C
H3C
O
O
HO
NH
NH
NodE, NodF
O
O
HO
OH
H 3C
O SO3H
OH
O
R6
NH
H3C
O
HO
O
NH
O
O
OH
R5
NH
reducing
terminus
H3C
H3C
OH
O
O
O
n
Acyl
R1
R2
R3
R4
R5
R6
n
C16, C18, C20 (varying degrees
of saturation; between 0 and 4
double bonds)
H,
Me
H, OH,
Cb
H, Ac,
Cb
H, S, Ac,
Fuc,
MeFuc,
AcFuc,
AcMeFuc,
SMeFuc
H,
Ara
H, OH,
Ac, Fuc
-1, 0,
1, 2, 3
Fig. 2 (a) Structure of Sinorhizobium meliloti Nod factor. The N-acetylglucosamine backbone is
synthesised by NodA, NodB and NodC; the acyl chain group is determined by Nod E and NodF;
O-sulfation is performed by NodH, NodP and NodQ; O-acetylation is performed by NodL.
(b) Generalised structure of naturally occurring Nod factors. Table shows major decorations and
variations in Nod factor structure based on species studied to date. Ac acetyl, Ara arabinosyl,
Cb carbamoyl, Fuc fucosyl, H hydrogen, Me methyl, OH hydroxyl, S sulfate, AcFuc acetylated
fucose, MeFuc methylfucose, AcMeFuc acetylated methylfucose, SMeFuc sulfated methylfucose.
Figure adapted from Perret et al. (2000) and Wais et al. (2002)
The Role of Diffusible Signals in the Establishment of Rhizobial and Mycorrhizal
7
Rhizobia also produce many other compounds (including type I and type III
secreted proteins and surface polysaccharides) which have varying degrees of
importance during nodulation. These will not be discussed here as Nod factors
are the most important diffusible signalling molecules when considering the establishment of the legume-Rhizobium symbiosis. A recent detailed review of these
additional signals is provided by Downie (2010).
2.2.1
Nod Factor Core (NodA, NodB, NodC)
The core structure of Nod factors is produced by the enzymes encoded for by the
Rhizobium genes nodABC, which form an operon in many species. The first step of
Nod factor biosynthesis involves the assembly of the chitin backbone through the
activity of NodC, an N-acetylglucosaminyltransferase, which causes chain elongation at the non-reducing terminus (Kamst et al. 1997, 1999; Mergaert et al. 1995;
Spaink et al. 1994). NodB, an N-deacetylase, then removes the N-acetyl moiety
from the non-reducing terminus (John et al. 1993; Spaink et al. 1994), allowing for
the subsequent addition of an acyl chain to the chitin oligomer via NodA, an
N-acyltransferase (Fig. 2a; Atkinson et al. 1994; Debelle et al. 1996; Rohrig et al.
1994). NodA represents a unique biosynthetic enzyme since it allows the addition
of an acyl chain to a polysaccharide without going through a nucleotide-activated
intermediate.
The nodABC genes are absolutely essential for Nod factor synthesis and bacterial mutants in these genes are unable to initiate plant signalling (Wais et al. 2002)
or nodulate their host (Fisher et al. 1985; Marvel et al. 1985). Cross-species
complementation experiments have demonstrated that nodABC loci of a number
of Rhizobium strains are able to complement mutants of a different strain and allow
successful nodulation (Krishnan and Pueppke 1991; Marvel et al. 1985). Therefore,
nodABC were previously referred to as the common nodulation genes. However,
recent sequencing of two Bradyrhizobium species (BTAi1 and ORS278) revealed
the absence of the nodABC genes from these genomes; despite this lack of nodABC,
Bradyrhizobium species ORS285 efficiently formed fully functional nodules
(Giraud et al. 2007). This exception to the rule therefore provides a unique example
of Nod factor-independent establishment of a root nodule symbiosis.
NodA and NodC have also been implicated in determining host-range specificity: S. meliloti NodA is able to transfer unsaturated C16 fatty acids to the Nacetylglucosamine backbone whilst R. tropici NodA is unable to perform this
reaction (Roche et al. 1996). Characterisation of NodC from different Rhizobium
species has demonstrated that this protein controls the number of N-acetylglucosamine residues which condense together: S. meliloti NodC forms chitotetraoses
whilst Mesorhizobium loti NodC forms chitopentaoses (Kamst et al. 1995, 1997).
However, it is the decorations added to the core Nod factor structure which play a
more important role in determining host-range specificity.
8
2.2.2
J.B. Miller and G.E.D. Oldroyd
Nod Factor Decorations
Research into Nod factor structural requirements began through a series of crossinoculation experiments whereby Rhizobium isolates from one legume were
inoculated onto other legume species to determine whether nodulation was possible. Although the precise structures of Nod factors had not been determined, these
experiments provided a wealth of data on host-range specificity. Now that many of
these Nod factor structures have been determined it is clear that the decorations
added to the core Nod factor structure (Fig. 2b) are important for determining hostrange specificity. For example, expression of nodABC in Escherichia coli is alone
sufficient to trigger root hair deformation in clover but not M. sativa; however, root
hair deformation is observed in both species upon the additional expression of nodH
(Banfalvi and Kondorosi 1989). Similar experiments have established roles for
other host-specific nod genes in controlling host-range specificity through the
decorations added to the core Nod factor structure (Lopez-Lara et al. 1996; Lorquin
et al. 1997a, b; Mergaert et al. 1996; Spaink et al. 1991). Additionally, this role for
different Nod factor structures in determining host-range specificity is particularly
important when considering the broad host-range Sinorhizobium sp. strain NGR234
which is able to nodulate many host plants because it produces a wide variety of
Nod factor structures (Price et al. 1992).
Acylation (NodE, NodF)
NodE and NodF determine which acyl chain(s) are added to the core Nod factor
structure by NodA. NodF is an acyl carrier protein, whilst NodE is a b-ketoacyl
synthase implicated in determining the degree of acyl chain saturation (Bloemberg
et al. 1995a; Debelle and Sharma 1986; Geiger et al. 1991; Ritsema et al. 1997; van
der Drift et al. 1996). In R. leguminosarum bv. viciae, NodE activity leads to the
production of a Nod factor with a polyunsaturated C18:4 acyl chain. Inactivation of
this gene instead results in the incorporation of vaccenic acid, an unsaturated C18:1
acyl chain (Spaink et al. 1991), and subsequently renders the strain unable to
nodulate Vicia sativa (Canter Cremers et al. 1989). Deletion of S. meliloti nodF
yields Nod factors with similar acyl chain compositions to those obtained from
nodE deletion mutants, suggesting that the combined action of NodE and NodF is
required for appropriate acyl chain addition (Demont et al. 1993). Indeed, exchanging the nodEF genes of S. meliloti with those of R. leguminosarum bv. viciae
extends the production of Nod factors to include structures with polyunsaturated
C18:2, C18:3 and C18:4 acyl chains (Demont et al. 1993). Methyl-branched acyl
chains are added to the Nod factor of the arctic Mesorhizobium sp. strain N33
(Oxytropis arctobia) and this requires a fully functional nodE gene (Poinsot et al.
2001). Poinsot et al. (2001) speculate that incorporation of these unusual acyl
chains is important for this species to tolerate extreme cold.
The Role of Diffusible Signals in the Establishment of Rhizobial and Mycorrhizal
9
Some species of legumes utilise specific recognition of the acyl chain as a
stringent measure of Nod factor during rhizobial infection (Ardourel et al. 1994;
Walker and Downie 2000). However, this recognition occurs in combination with
either the O-acetylation of Nod factor by NodL (Sect. 2.2.2.3; Ardourel et al. 1994),
or the action of other Nod proteins (Walker and Downie 2000).
Glycosylation (NoeC, NodZ, NolK)
Two forms of glycosylation have been described as Nod factor decorations:
arabinosylation and fucosylation. The importance of both modifications has been
described for Nod factors from Azorhizobium caulinodans, the symbiont of
Sesbania rostrata (Mergaert et al. 1997). Arabinosylation and fucosylation also
appear to be required by other symbionts of S. rostrata, suggesting a possible
responsibility of these glycosyl decorations for determining host-range specificity
(Lorquin et al. 1997a).
D-arabinosylation on C3 of the reducing terminus of Nod factor is dependent on
noeC and/or downstream genes in A. caulinodans (Mergaert et al. 1996). Presence
of this D-arabinosyl group on Nod factors from A. caulinodans results in higher
numbers of nodules on S. rostrata roots than Nod factors without this decoration
(Fernandez-Lopez et al. 1998). However, other host species of A. caulinodans show
a preference for fucosylated Nod factors, implying that arabinosylation is particularly important for nodulation of S. rostrata (Fernandez-Lopez et al. 1998).
L-fucosylation of A. caulinodans Nod factors on C6 of the reducing terminus
depends on both nodZ and nolK. NolK is involved in the biosynthesis of GDPfucose, which is a substrate for the fucosyltransferase NodZ (Mergaert et al. 1996).
These authors also detected a NodZ-independent Nod factor fucosyltransferase
activity, although this activity was not encoded for by any of the known nod genes.
NodZ also decorates the Nod factors of other Rhizobium species with fucosyl
groups (Lopez-Lara et al. 1996; Quesada-Vincens et al. 1997; Quinto et al. 1997;
Stacey et al. 1994). NodZ from Sinorhizobium sp. strain NGR234 preferentially
fucosylates chitopentaoses over single N-acetylglucosamine residues or nonfucosylated Nod factors, implying that fucosylation occurs before acylation
(Quesada-Vincens et al. 1997). Nodulation of Macroptilium atropurpureum by
B. japonicum (Stacey et al. 1994) and Pachyrhizus tuberosus by Sinorhizobium
sp. strain NGR234 (Quesada-Vincens et al. 1997) is blocked by mutation of nodZ.
However, expression of B. japonicum nodZ in R. leguminosarum bv. viciae results
in fucosylated Nod factor production and extends the host range of the
R. leguminosarum strain (Lopez-Lara et al. 1996). It is also interesting to note
that despite the absolute necessity of B. japonicum for NodZ in order to nodulate
M. atropurpureum, nodZ expression is not controlled by NodD, a trait which is
unique amongst the nod genes (Stacey et al. 1994).
An additional rare fucosylation site has been identified in Mesorhizobium loti
strain NZP2213 where the fucosyl residue is found on a non-terminal N-acetylglucosamine residue of the Nod factor structure (Olsthoorn et al. 1998).
10
J.B. Miller and G.E.D. Oldroyd
Acetylation (NodL, NodX, NolL)
An O-acetyl group can be added on C6 of either the reducing or non-reducing
terminal N-acetylglucosamine residue of Nod factor through the action of NodX
or NodL, respectively (Bloemberg et al. 1994). NodX from R. leguminosarum
bv. viciae is able to only use chitopentaoses as a substrate for O-acetylation
(Firmin et al. 1993) and the quantities of O-acetylated Nod factor produced by
NodX is temperature-dependent (Olsthoorn et al. 2000). Mutation of nodX in
R. leguminosarum bv. viciae strain TOM abolishes the ability of this strain to
nodulate Pisum sativum cv. Afghanistan (Davis et al. 1988). This nodX mutant
produces other LCOs identical to wild-type bacteria (Ovtsyna et al. 1999),
suggesting that the specificity of the interaction between strain TOM and P. sativum
cv. Afghanistan is due to O-acetylation. However, nodulation with this nodX
mutant can be restored by expression of nodZ from B. japonicum (Sect. 2.2.2.2;
Ovtsyna et al. 1998), implying that O-acetylation alone cannot be the only mechanism for determining specificity in this interaction.
The NodL O-acetylation reaction is dependent upon a non-reducing terminally
de-N-acetylated chitin oligosaccharide substrate (i.e. the product of NodB and
NodC activity; Bloemberg et al. 1995b). The stringency for NodL-mediated Oacetylation appears to be low since fully functional nodulation of M. sativa by the
S. meliloti nodL mutant is possible, although significantly decreased infection
thread formation and a delay in nodulation was noted (Ardourel et al. 1994).
In addition to acetylation by NodX or NodL, acetyl groups can be added onto
fucose decorations via NolL. NolL from Sinorhizobium sp. strain NGR234 leads to
Nod factor structures with 3-O- or 4-O-acetylation on fucose (Berck et al. 1999),
whilst NolL from Rhizobium etli yields Nod factors with only 4-O-acetylation on
fucose (Corvera et al. 1999). NolL is not essential for nodulation of Phaseolus
vulgaris by R. etli, although nodulation of the nolL mutant was less efficient than
the wild-type strain on some P. vulgaris cultivars (Corvera et al. 1999). Heterologous expression of nodZ or nodZ and nolL has demonstrated that NolL is necessary
for efficient nodulation of Lotus japonicus by R. leguminosarum bv. viciae (Pacios
Bras et al. 2000). Different Lotus species also have different requirements for NolLmediated acetylation on fucose: the M. loti nolL mutant is unable to form infected
nodule primordia on L. filicaulis and L. corniculatus yet can successfully nodulate
L. japonicus (Rodpothong et al. 2009). This apparent discrepancy for the requirement of NolL for successful nodulation of L. japonicus was explained by
Rodpothong et al. (2009) as being due to other differences, notably the acyl chain
structure, between the Nod factors of M. loti (the true symbiont of L. japonicus) and
R. leguminosarum bv. viciae expressing nodZ and nolL.
A rare acetylation site has been determined in the M. loti strain N33 where 6-Oacetylation occurs on the residue proximal to the non-reducing N-acetylglucosamine, although the gene encoding the enzyme responsible for this modification
has not been identified (Poinsot et al. 2001).
The Role of Diffusible Signals in the Establishment of Rhizobial and Mycorrhizal
11
Methylation (NodS, NoeI)
Two forms of methylation can occur on Nod factors: N-methylation on the nonreducing terminus controlled by NodS (Geelen et al. 1993; Jabbouri et al. 1995) or
2-O-methylation on fucose mediated by NoeI (Jabbouri et al. 1998). NodS is an Nmethyltransferase and in A. caulinodans or Sinorhizobium sp. strain NGR234
methylates end-deacetylated chitooligosaccharides using an S-adenosyl-L-methionine-binding protein as a methyl donor (Geelen et al. 1995). Indeed, the structure of
NodS from B. japonicum has recently been solved, representing the first crystal
structure of an S-adenosyl-L-methionine-dependent methyltransferase (Cakici et al.
2010). The R. etli nodS mutant is less able to induce root hair curling and actin
cytoskeleton rearrangements in P. vulgaris than wild-type R. etli, suggesting that
N-methylation is key in regulating these Nod factor-dependent responses (Cardenas
et al. 2003). N-methylation by NodS biosynthetically precedes any O-acetylation
reactions by NodL (Lopez-Lara et al. 2001).
2-O-methylation of fucose by NoeI is common in Sinorhizobium sp. strain
NGR234 and S. fredii strain USDA257 (Jabbouri et al. 1998). Mutation of this
gene leads to production of LCOs which are non-methylation on fucose, although as
this appears to have no effect on nodulation (Jabbouri et al. 1998) this Nod factor
decoration is of lesser significance.
Carbamoylation (NodU, NolO)
Carbamoylation on the non-reducing terminal N-acetylglucosamine residue is controlled by the carbamoyltransferases NodU and NolO. Expression of Sinorhizobium
sp. strain NGR234 nodU in S. fredii strain USDA257 (which does not produce
carbamoylated Nod factors) allows 6-O-carbamoylated Nod factors production
(Jabbouri et al. 1995). Likewise, expression of Sinorhizobium sp. strain NGR234
nolO in S. fredii strain USDA257 has confirmed the role of NolO in controlling 3O- and 4-O-carbamoylation at the non-reducing terminus (Jabbouri et al. 1998). The
host range of S. fredii expressing nolO is increased to include non-host species
(Jabbouri et al. 1998). However, the nodulation phenotype of the S. fredii nolO
mutant was not different from the wild-type strain, although the mutant showed
decreased competitiveness to nodulate G. max (Madinabeitia et al. 2002). Interestingly, Jabbouri et al. (1998) suggest the existence of a third (as yet uncharacterised)
carbamoyltransferase in Sinorhizobium sp. strain NGR234 since mutation of nodU
and nolO failed to result in Nod factors entirely devoid of carbamoylation.
Sulfation (NodH, NodP, NodQ, NoeE)
Addition of an O-sulfate group is common to many Nod factors and this reaction is
performed by the sulfotransferases NodH (Del Papa et al. 2007; Ehrhardt et al.
1995; Laeremans et al. 1996; Lerouge et al. 1990; Roche et al. 1991; Schultze et al.
12
J.B. Miller and G.E.D. Oldroyd
1995) and NoeE (Hanin et al. 1997). NodH activity results in sulfation on C6 of the
reducing terminus, while NoeE only gives sulfation on fucose residues attached to
C6 of the reducing terminus (Quesada-Vincens et al. 1998). NodP and NodQ are
also essential for Nod factor sulfation and act as sulfur activators by synthesising
the sulfur donor 30 -phosphoadenosine 50 -phosphosulfate (Schwedock and Long
1990; Schwedock et al. 1994).
The R. tropici strain CFN299 nodP mutant shows decreased nodulation on
P. vulgaris cv. Negro Xamapa, while nodH and nodP mutants acquire an increased
capacity to nodulate the two other cultivars (Laeremans et al. 1996). Likewise, the
nodH mutant of R. tropici shows decreased nodulation in comparison to wild-type
when nodulating Leucaena leucocephala (Folch-Mallol et al. 1996). R. fredii
expressing noeE produced sulfated LCOs and therefore acquired the ability to
nodulate Calopogonium caeruleum, whilst mutation of noeE from Sinorhizobium
sp. strain NGR234 abolished the production of sulfated LCOs and prevented
nodulation of P. tuberosus (Hanin et al. 1997). Importantly, S. meliloti Nod factor
sulfation is essential for root hair deformation and nodulation of M. sativa (Roche
et al. 1991). These findings all support a role for sulfation as a major determinant of
symbiont specificity for their host plant species, yet in other interactions the
stringency for sulfation appears to be low. Mutation of the nodHPQ genes of
Rhizobium sp. strain N33 appears to have no effect on nodulation of two host
species tested (Cloutler et al. 1996). Remarkably, a recent report suggests that the
nodH mutant of Sinorhizobium sp. strain BR816 shows increased nitrogen fixation
relative to the wild-type strain despite there being no other nodulation phenotype
(Remans et al. 2007). The authors attribute this interesting result to the availability
of activated sulfate inside nodules which they argue is likely to be greater with the
nodH mutant.
2.2.3
Nod Factor Secretion
NodI and NodJ act as an ATP-binding cassette (ABC) transporter (Higgins et al.
1986) and are involved in Nod factor secretion. Secretion of Nod factor is impaired
in nodI and/or nodJ rhizobia mutants (Cardenas et al. 1996; Fernandez-Lopez et al.
1996; Spaink et al. 1995), whilst E. coli engineered for the biosynthesis of Nod
factors only secreted these LCOs in the presence of NodI and NodJ (FernandezLopez et al. 1996). Interestingly, the nodIJ mutant of R. etli is able to nodulate P.
vulgaris, although the mutant shows a delayed and decreased nodulation phenotype
in comparison to the wild-type. This non-essential role of NodI and NodJ therefore
suggests a possible additional component involved in rhizobial secretion of Nod
factors which has yet to be characterised.
The Role of Diffusible Signals in the Establishment of Rhizobial and Mycorrhizal
2.3
13
Plant Responses to Nod Factor
Nod factors are perceived in epidermal and root hair cells since fluorescent Nod
factors added to plant roots accumulate in the walls of these cells (Goedhart et al.
2000). Host plant cells are able to perceive Nod factor concentrations as low as
1012 M (Oldroyd and Downie 2004), suggesting that the receptor able to perceive
Nod factors is highly sensitive. In L. japonicus, two Nod factor receptors have been
identified: NFR1 and NFR5 (Madsen et al. 2003; Radutoiu et al. 2003). The M.
truncatula gene equivalent to NFR5 is NFP (Nod factor perception), whilst LYK3 is
orthologous to NFR1 (Amor et al. 2003; Smit et al. 2007). Interestingly, LYK3 has
been described as an entry receptor which controls rhizobial infection in a manner
dependent upon Nod factor structure (Smit et al. 2007), therefore suggesting an
additional element for Nod factor structural specificity in establishing the legumeRhizobium symbiosis. These Nod factor receptors are receptor-like kinases and
contain LysM domains which are involved in binding N-acetylglucosamine,
making them likely Nod factor receptors. Importantly, M. truncatula transformed
with NFR1 and/or NFR5 was able to form nodules with a rhizobial species usually
specific to L. japonicus (Radutoiu et al. 2007). This directly implicates NFR1 and
NFR5 as Nod factor receptors whilst also demonstrating the importance of these
receptors and the structure of Nod factors themselves for determining symbiont
specificity. However, binding assays between Nod factors and their potential targets
have yet to provide formal evidence for a direct physical interaction between ligand
and receptor.
Nod factors trigger a range of molecular responses in legumes (reviewed by
D’Haeze and Holsters 2002; Oldroyd et al. 2001a). These responses include rapid
pH changes (Felle et al. 1996, 2000), root hair deformation (Roche et al. 1991;
Spaink et al. 1991), lateral root formation (Olah et al. 2005), reactive oxygen
species production (Cardenas et al. 2008; Cardenas and Quinto 2008), induction
of calcium flux (Ehrhardt et al. 1992), induction of calcium spiking (Ehrhardt et al.
1996), and gene expression changes (Mitra et al. 2004). Nod factors are required for
infection thread development but are insufficient to activate this response alone
(Dazzo et al. 1991), although the formation of pre-infection thread structures has
been described (van Brussel et al. 1992). At sufficiently high concentrations Nod
factors can also induce cortical cell division and the formation of nodule primordia
(Truchet et al. 1991). The diversity of these responses only goes to demonstrate the
critical importance of Nod factors as signalling molecules in the early stages of
nodulation.
The sensitivity of the responses triggered by Nod factor can vary by several
orders of magnitude; for example, the two Ca2+ signatures (flux and spiking) can be
separated, such that high concentrations (>109 M) of Nod factor induce flux
followed by spiking, while low Nod factor concentrations (<1010 M) induce
only calcium spiking (Shaw and Long 2003). Pisum sativum plants treated with
chitin oligomers of four or five residues also show calcium spiking, but not calcium
flux (Walker et al. 2000). These observations suggest that the activation of calcium
14
J.B. Miller and G.E.D. Oldroyd
spiking has a lower stringency for Nod factor structure and concentration than the
induction of calcium flux. The structural requirements of S. meliloti Nod factors to
trigger calcium spiking have been analysed and effects due to missing decorations,
such as O-acetylation, N-acylation or O-sulfation, can be overcome by treating with
high enough concentrations of Nod factor (Oldroyd et al. 2001b; Wais et al. 2002).
This work has formally shown that Nod factor decorations, in addition to determining host-range specificity, play a role in determining the potency of the Nod factor
signal to the plant and that concentration of LCOs must therefore be considered
when determining biological activity.
Calcium spiking, the rapid oscillation of calcium concentration in the nucleus
and peri-nuclear region of root hair cells (Ehrhardt et al. 1996), is central to the
signalling pathway activated by Nod factor (Oldroyd and Downie 2004).
Components of this common symbiosis signalling pathway are required for both
nodulation and mycorrhization in legumes; mutation in any of these genes blocks
the formation of either symbiosis.
3 Diffusible Signals During Mycorrhization
The molecular dialogue between AM fungi and host plant roots has been less well
characterised than that of legumes and rhizobia, partly because the fungus is an
obligate biotroph which makes it less amenable to study. An additional problem for
the study of AM fungi is the asynchronous nature of the infection process. Despite
these limitations, research in this area has begun to provide some interesting
parallels between signalling during nodulation and mycorrhization (reviewed by
Bonfante and Genre 2010; Harrison 2005; Parniske 2008). Diffusible signals again
play a key role in the establishment of mycorrhizal interactions. Strigolactones are
released from plant roots and these promote AM fungal spores to germinate.
Germinated spores produce a diffusible signal, a so-called “Myc” factor, which
triggers signalling in the plant (mediated by the symbiosis signalling pathway in
both legumes and non-legumes; Gutjahr et al. 2008). The chemical nature of “Myc”
factor has proved elusive, although recent work has characterised LCOs derived
from AM fungi which act in an analogous fashion to Nod factor.
3.1
Strigolactones
Strigolactones play an important role in establishing mycorrhizal symbioses,
serving as germination and hyphal branching cues for dormant AM fungal spores.
Nothing is known about strigolactone perception by the fungus or the requirement
for different chemical structures of strigolactones. Other diffusible signals, including flavonoids, have also been implicated in triggering spore germination.
The Role of Diffusible Signals in the Establishment of Rhizobial and Mycorrhizal
3.1.1
15
Structure and Synthesis
The first strigolactone isolated from root exudates was strigol (Cook et al. 1966).
Numerous subsequent experiments have demonstrated the presence of
strigolactones in root exudates from different species, including both monocotyledonous (Awad et al. 2006) and dicotyledonous plants (Yoneyama et al. 2008). It
was not until 2003 though that strigolactones were formally proved to be derived
from roots, as demonstrated through aseptic plant culture experiments (Yasuda
et al. 2003). Many chemical structures of naturally occurring strigolactones have
now been proposed (Yoneyama et al. 2009); these vary primarily in the position and
number of hydroxyl, methyl and acetyl groups present on the core ring structure.
Interestingly, Arabidopsis thaliana, a non-mycorrhizal species, produces low
concentrations of strigolactones relative to other plant species which can form
symbioses with AM fungi (Westwood 2000).
Relatively little is known about the biosynthesis of strigolactones, which were
originally considered to be a group of sesquiterpene lactones. However, the tricyclic ring structure of strigolactones is now known to be derived from carotenoid
biosynthesis (Fig. 3; Jamil et al. 2010; Lopez-Raez et al. 2008; Matusova et al.
2005). Two proteins characterised in A. thaliana as carotenoid cleavage
dioxygenase enzymes have been implicated specifically in the biosynthesis of
strigolactones: CCD7 (Booker et al. 2004) and CCD8 (Sorefan et al. 2003).
CCD7 and CCD8 were originally studied for their mutant phenotypes in shoot
branching and a role for branching inhibition by strigolactones is now established
(Gomez-Roldan et al. 2008; Umehara et al. 2008). Mutation or silencing which
causes decreased CCD7 expression in tomato plants gives rise to strigolactoneimpaired lines which show decreased colonisation with AM fungi, thus
demonstrating the importance of this enzyme for the synthesis of strigolactones
and the importance of strigolactones for mycorrhization (Koltai et al. 2010; Vogel
et al. 2010).
3.1.2
Biological Activity
Strigolactones have been well characterised for their role in the interaction between
plants and weeds of the genus Striga, from where these molecules derive their
name. During this parasitic interaction, strigolactones released by the host plant
promote germination of Striga species (reviewed by Bouwmeester et al. 2003).
However, the role of strigolactones during symbiotic interactions with AM fungi
was not determined until 2005 (Akiyama et al. 2005).
The amount of strigolactone secretion by roots is thought to be very low, but
these molecules are highly potent and are able to induce fungal hyphal branching at
picogram to nanogram quantities (Akiyama and Hayashi 2006; Bucher et al. 2009).
For this reason, synthetic strigolactones have also been used in the study of
strigolactones and AM fungi. Fungal responses after the application of synthetic
16
J.B. Miller and G.E.D. Oldroyd
Pyruvate + Glyceraldehyde-3-phosphate
2-methylerythritol-4-phosphate (MEP)
Isopentenyl diphosphate (IPP)
Dimethylallyl pyrophosphate
IPP isomerase
O
O
GGPP synthase
Geranyl pyrophosphate (C10)
O
O
Synthetic
strigolactone (GR24)
O
GGPP synthase
IPP
Farnesyl pyrophosphate (C15)
CH3
R2
R3
O
O
Natural
strigolactones
DITERPENES
Phytoene synthase
GGPP
O
SESQUITERPENES
GGPP synthase
IPP
Geranylgeranyl pyrophosphate (GGPP)(C20)
R1 CH3
MONOTERPENES
Phytoene (C40)
O
O
CH3
Lycopene
a -carotene
CCD7
CCD8
b-carotene
STRIGOLACTONES
ABA
Fig. 3 Biosynthesis of strigolactones. Partial diagram of the carotenoid biosynthesis pathway.
Enzymes are depicted in italics; the 2-methylerythritol-4-phosphate (MEP) pathway and the first
committed steps of carotenoid biosynthesis are denoted by upper and lower shaded boxes,
respectively; compounds in dashed boxes represent major side branches of the pathway. ABA
abscisic acid, CCD7/8 carotenoid cleavage dioxygenase, GGPP geranylgeranyl pyrophosphate,
IPP isopentenyl diphosphate. Insets show chemical structures of example synthetic and naturally
occurring strigolactones: strigol: R1 ¼ CH3, R2 ¼ OH, R3 ¼ H; strigyl acetate: R1 ¼ CH3, R2 ¼
OAc, R3 ¼ H; sorgolactone: R1 ¼ H, R2 ¼ H, R3 ¼ H. Figure adapted from Akiyama and
Hayashi (2006) and Matusova et al. (2005)
strigolactones, such as GR24, are comparable to treatment with naturally occurring
strigolactones (Akiyama et al. 2005). Strigolactones (or “branching factors” as they
were originally named before their identification) promote spore germination and
branching in Gigaspora spp. (Akiyama et al. 2005; Buee et al. 2000; Nagahashi and
Douds 1999). In addition, strigolactone treatment promotes a number of other
fungal pre-symbiotic responses, including the induction of mitosis (Buee et al.
2000), increased expression of mitochondrial-related genes (Tamasloukht et al.
2003), increased density of mitochondria (Besserer et al. 2009), and thus increased
respiratory activity (Tamasloukht et al. 2003).
Mycorrhization is impaired in the strigolactone-deficient ccd8 mutant of
P. sativum, although this phenotype can be partially recovered by exogenous
application of GR24 (Gomez-Roldan et al. 2008). As full complementation is not
achieved with exogenous treatment of strigolactone it is possible that directionality
The Role of Diffusible Signals in the Establishment of Rhizobial and Mycorrhizal
17
of the diffusible signal via a concentration gradient is important in order to
encourage germinating AM spores to grow towards host roots. This existence and
importance of a natural concentration gradient is especially likely when considering
that strigolactones are readily hydrolysed in the soil (Akiyama and Hayashi 2006).
Evidence for chemotaxis responses of AM fungi to root diffusible signals supports
this hypothesis (Sbrana and Giovannetti 2005).
The involvement of flavonoids as diffusible signals during the establishment of
interactions with AM fungi remains unclear (as discussed by Larose et al. 2002;
Vierheilig et al. 1998). For example, the flavonoid medicarpin accumulates in
M. sativa roots and strongly inhibits Glomus intraradices hyphal growth
(Guenoune et al. 2001), whilst a flavonoid from melon roots enhances
mycorrhization in this species (Akiyama et al. 2002). Likewise, the flavonoid
quercetin stimulates AM fungal spore growth and branching (Becard et al. 1992;
Tsai and Phillips 1991). Other studies have suggested that flavonoids are not
absolutely essential for hyphal growth (Becard et al. 1995). The most likely
conclusion is that AM fungal responses to flavonoids are compound and genus
specific (Scervino et al. 2005a, b), therefore making it difficult to assign a definitive
role for these diffusible signals during the establishment of symbioses with AM
fungi. Contrasting this with nodulation, recent evidence suggests that nodulation of
M. sativa by S. meliloti is increased by strigolactone treatment (Soto et al. 2010); it
will therefore be interesting to know whether the interplay between these diffusible
signals is important for the establishment of both symbioses.
3.2
“Myc” Factors
A diffusible signal originating from the fungus, the so-called “Myc” factor, has long
been hypothesised, but until recently had not been characterised. It has been shown
that a diffusible factor released from germinating AM fungi is able to induce
expression of ENOD11, a symbiosis-specific gene in M. truncatula (Kosuta et al.
2003). Olah et al. (2005) used a membrane separating AM fungi from plants to
demonstrate that this diffusible fungal factor identified by Kosuta et al. (2003)
activates root branching in M. truncatula (a response also observed with Nod
factor). AM fungi are able to induce calcium spiking in M. truncatula root hair
cells associated with highly branched fungal hyphae and this occurs prior to
physical contact between the fungus and the root (Kosuta et al. 2008). Calcium
spiking has also been detected upon contact of fungal hyphopodia with nontrichoblastic root cells of M. truncatula and Daucus carota, and also upon treatment
of M. truncatula roots with a concentrated extract from germinating fungal spores
(Chabaud et al. 2011). This Ca2+ spiking in M. truncatula is dependent on
components of the symbiosis signalling pathway but is NFP-(in)dependent, therefore implying different plant machineries for the recognition of “Myc” and Nod
factors (Chabaud et al. 2011; Kosuta et al. 2008). Calcium transients have also been
detected in G. max cell cultures exposed to germinating spores from Glomus species
18
J.B. Miller and G.E.D. Oldroyd
(Navazio et al. 2007), although this signal was also released by non-germinating
spores so perhaps represents a triggering of defence responses.
3.2.1
LCOs as a “Myc” Factor
An exciting recent development has proposed the structures of two LCOs produced
by the AM fungus G. intraradices (Fig. 4; Maillet et al. 2011). One of these LCOs
contains an unsaturated C18:1 acyl chain and is non-sulfated on the reducing
terminus (Fig. 4a), whilst the other LCO has a saturated C16 acyl chain and is Osulfated at the reducing terminus (Fig. 4b). These LCOs were characterised for their
ability to induce pENOD11::GUS expression and root hair deformation in M.
truncatula. Application of these Myc LCOs to M. truncatula, Tagetes patula or
D. carota resulted in increased mycorrhizal colonisation. Increased lateral root
formation in M. truncatula was also observed upon Myc LCO treatment and
importantly this was NFP-dependent (Maillet et al. 2011). It has been previously
a
O
HO
HO
NH
O
O
HO
OH
OH
OH
OH
O
O
O
HO
NH
H3C
O
O
HO
OH
NH
NH
H3C
H3C
O
O
O
b
O
HO
HO
O
O
HO
NH
NH
H3C
O
O SO3H
OH
OH
OH
O
HO
O
NH
H3C
OH
NH
H3C
O
O
O
O
O
HO
CH3
CH3
c
O
HO
HO
NH
Acyl
O
HO
O R
OH
OH
OH
O
O
HO
O
O
HO
H3C
H3C
O
O
OH
NH
NH
NH
H3C
O
n
n
O
Acyl
R
n
C16:0, C16:1,
C16:2, C18:0,
C18:1
H,
S
1,
2
Fig. 4 (a) Structure of Glomus intraradices non-sulfated lipochitooligosaccharide (LCO). (b)
Structure of G. intraradices sulfated LCO. (c) Generalised structure of naturally occurring G.
intraradices LCOs. Table shows decorations and variations in Myc LCOs. Abbreviations as in
Fig. 2. Figure adapted from Maillet et al. (2011)
The Role of Diffusible Signals in the Establishment of Rhizobial and Mycorrhizal
19
shown that NFP is not required for mycorrhizal associations (Amor et al. 2003);
therefore the apparent discrepancy for the requirement of NFP for inducing different mycorrhizal responses in M. truncatula could be due to the existence of other as
yet uncharacterised diffusible signals derived from AM fungi. The NFP-independent diffusible fungal signals described by Olah et al. (2005) and Chabaud et al.
(2011) may therefore represent different novel classes of diffusible signalling
molecules (i.e. non-LCOs) involved in establishing the mycorrhizal symbioses.
The apparent dual role of NFP in signalling during mycorrhization and nodulation
could also be explained by the existence of a receptor complex consisting of NFP
and other currently uncharacterised receptors. This is supported by the dual role of
Parasponia NFP, which is involved in both nodulation and mycorrhization (Op den
Camp et al. 2011). Alternatively, if early mycorrhizal signalling occurs in a directly
analogous fashion to that of Nod factor, the “Myc” factor which causes NFPindependent root branching (Olah et al. 2005) and calcium spiking (Chabaud
et al. 2011; Kosuta et al. 2008) would require an additional separate receptor.
However, such a “Myc” factor receptor has yet to be identified.
Given the diversity of Nod factor structures and the fact that at least 80% of land
plants are able to engage in symbiotic interactions with mycorrhiza, it is almost
certain that other Myc LCOs exist in nature and have yet to be isolated and
characterised. It is also tempting to speculate that G. intraradices and other AM
fungi produce a broad spectrum of LCOs in order to colonise a wide range of host
species, as with the broad host-range Sinorhizobium sp. strain NGR234.
4 Conclusions and Perspectives
The importance of diverse diffusible signals during the establishment of nodulation
and mycorrhization is clear. Nod factor structural diversity has been implicated as a
key determinate of host-range specificity, as discussed here and reviewed extensively by D’Haeze and Holsters (2002) and Perret et al. (2000). However, this
structural diversity alone does not determine host-range specificity in all legumeRhizobium symbioses; for example, R. etli and M. loti produce identical Nod factors
yet nodulate different plant species (Cardenas et al. 1995). The recent identification
of LCOs produced by the AM fungus G. intraradices will no doubt result in detailed
research into the structural importance of these molecules during mycorrhizal
interactions. Developments in signalling by nodulation- and mycorrhization-specific LCOs will also prove exciting, particularly in addressing the question of
specificity in symbiosis signalling.
When considering the signals released from symbionts it is important to bear in
mind that mycorrhization evolved ~400 million years before nodulation (Kistner
and Parniske 2002). Chitin, the major component of fungal cell walls, therefore
becomes a key molecule: chitin fragments released from cell walls of symbiotic or
pathogenic fungi would act as a trigger of plant defence responses. In order to
differentiate themselves as symbiotic (and also to protect against hydrolysis by
20
J.B. Miller and G.E.D. Oldroyd
plant-released chitinases) it would be essential that AM fungi modify these
structures with decorations. For rhizobia to adopt fungal genes responsible for
Myc LCO production and thereby gain the ability to synthesise LCOs becomes a
tangible explanation for the existence of Nod factors. It will therefore be interesting
whether the genomes of AM fungi contain homologues to any rhizobial nod genes.
The question then turns to pathogens and why these too have not evolved diffusible
signals such as LCOs in order to hijack symbiotic signalling. Since plants regulate
their symbioses by monitoring symbiotic efficiency (Schumpp and Deakin 2010) it
is plausible that pathogenic microorganisms which do abuse this symbiotic molecular dialogue and gain entry to the root are detected and killed by the host species.
Indeed, the combined action of diffusible LCOs and many additional non-diffusible
signals is essential in order to discriminate between pathogenic and symbiotic
microorganisms (Hamel and Beaudoin 2010).
In conclusion, diffusible signals from both host plant and symbiont are essential
for the establishment of nodulation and mycorrhization. Work since the original
characterisation of Nod factors has irrefutably demonstrated the importance of
these rhizobial-derived LCOs during the legume-Rhizobium symbiosis. Future
research into mycorrhizal-derived LCOs will no doubt identify novel LCO
structures produced by AM fungi and hopefully begin to shed light on the
mechanisms of discriminating specificity during symbiosis signalling.
References
Akiyama K, Matsuoka H, Hayashi H (2002) Isolation and identification of a phosphate deficiencyinduced C-glycosylflavonoid that stimulates arbuscular mycorrhiza formation in melon roots.
Mol Plant Microbe Interact 15:334–340
Akiyama K, Matsuzaki K, Hayashi H (2005) Plant sesquiterpenes induce hyphal branching in
arbuscular mycorrhizal fungi. Nature 435:824–827
Akiyama K, Hayashi H (2006) Strigolactones: chemical signals for fungal symbionts and parasitic
weeds in plant roots. Ann Bot 97:925–931
Amor BB, Shaw SL, Oldroyd GE, Maillet F, Penmetsa RV, Cook D, Long SR, Denarie J, Gough C
(2003) The NFP locus of Medicago truncatula controls an early step of Nod factor signal
transduction upstream of a rapid calcium flux and root hair deformation. Plant J 34:495–506
Ardourel M, Demont N, Debelle F, Maillet F, de Billy F, Prome JC, Denarie J, Truchet G (1994)
Rhizobium meliloti lipooligosaccharide nodulation factors: different structural requirements
for bacterial entry into target root hair cells and induction of plant symbiotic developmental
responses. Plant Cell 6:1357–1374
Atkinson EM, Palcic MM, Hindsgaul O, Long SR (1994) Biosynthesis of Rhizobium meliloti
lipooligosaccharide Nod factors: NodA is required for an N-acyltransferase activity. Proc Natl
Acad Sci USA 91:8418–8422
Awad A, Sato D, Kusumoto D, Kamioka H, Takeuchi Y, Yoneyama K (2006) Characterization of
strigolactones, germination stimulants for the root parasitic plants Striga and Orobanche
produced by maize, millet and sorghum. Plant Growth Regul 48:221–227
Banfalvi Z, Kondorosi A (1989) Production of root hair deformation factors by Rhizobium meliloti
nodulation genes in Escherichia coli: HsnD (NodH) is involved in the plant host-specific
modification of the NodABC factor. Plant Mol Biol 13:1–12
The Role of Diffusible Signals in the Establishment of Rhizobial and Mycorrhizal
21
Becard G, Douds DD, Pfeffer PE (1992) Extensive in vitro hyphal growth of vesicular-arbuscular
mycorrhizal fungi in the presence of CO(2) and flavonols. Appl Environ Microbiol 58:821–825
Becard G, Taylor LP, Douds DD, Pfeffer PE, Doner LW (1995) Flavonoids are not necessary plant
signal compounds in arbuscular mycorrhizal symbioses. Mol Plant Microbe Interact 8:252–258
Berck S, Perret X, Quesada-Vincens D, Prome J, Broughton WJ, Jabbouri S (1999) NolL of
Rhizobium sp. strain NGR234 is required for O-acetyltransferase activity. J Bacteriol
181:957–964
Besserer A, Becard G, Roux C, Sejalon-Delmas N (2009) Role of mitochondria in the response of
arbuscular mycorrhizal fungi to strigolactones. Plant Signal Behav 4:75–77
Bloemberg GV, Thomas-Oates JE, Lugtenberg BJ, Spaink HP (1994) Nodulation protein NodL of
Rhizobium leguminosarum O-acetylates lipo-oligosaccharides, chitin fragments and
N-acetylglucosamine in vitro. Mol Microbiol 11:793–804
Bloemberg GV, Kamst E, Harteveld M, van der Drift KM, Haverkamp J, Thomas-Oates JE,
Lugtenberg BJ, Spaink HP (1995a) A central domain of Rhizobium NodE protein mediates host
specificity by determining the hydrophobicity of fatty acyl moieties of nodulation factors. Mol
Microbiol 16:1123–1136
Bloemberg GV, Lagas RM, van Leeuwen S, Van der Marel GA, Van Boom JH, Lugtenberg BJ,
Spaink HP (1995b) Substrate specificity and kinetic studies of nodulation protein NodL of
Rhizobium leguminosarum. Biochemistry 34:12712–12720
Bonfante P, Genre A (2010) Mechanisms underlying beneficial plant-fungus interactions in
mycorrhizal symbiosis. Nat Commun 1:1–11
Booker J, Auldridge M, Wills S, McCarty D, Klee H, Leyser O (2004) MAX3/CCD7 is a
carotenoid cleavage dioxygenase required for the synthesis of a novel plant signaling molecule.
Curr Biol 14:1232–1238
Bouwmeester HJ, Matusova R, Zhongkui S, Beale MH (2003) Secondary metabolite signalling in
host-parasitic plant interactions. Curr Opin Plant Biol 6:358–364
Brachmann A, Parniske M (2006) The most widespread symbiosis on Earth. PLoS Biol 4:e239
Brencic A, Winans SC (2005) Detection of and response to signals involved in host-microbe
interactions by plant-associated bacteria. Microbiol Mol Biol Rev 69:155–194
Broughton WJ, Jabbouri S, Perret X (2000) Keys to symbiotic harmony. J Bacteriol 182:
5641–5652
Bucher M, Wegmuller S, Drissner D (2009) Chasing the structures of small molecules in
arbuscular mycorrhizal signaling. Curr Opin Plant Biol 12:500–507
Buee M, Rossignol M, Jauneau A, Ranjeva R, Becard G (2000) The pre-symbiotic growth of
arbuscular mycorrhizal fungi is induced by a branching factor partially purified from plant root
exudates. Mol Plant Microbe Interact 13:693–698
Burn JE, Rossen L, Johnston AW (1987) Four classes of mutations in the nodD gene of Rhizobium
leguminosarum biovar. viciae that affect its ability to autoregulate and/or activate other nod
genes in the presence of flavonoid inducers. Genes Dev 1:456–464
Caetano-Anolles G, Crist-Estes DK, Bauer WD (1988) Chemotaxis of Rhizobium meliloti to the
plant flavone luteolin requires functional nodulation genes. J Bacteriol 170:3164–3169
Cakici O, Sikorski M, Stepkowski T, Bujacz G, Jaskolski M (2010) Crystal structures of NodS Nmethyltransferase from Bradyrhizobium japonicum in ligand-free form and as SAH complex.
J Mol Biol 404:874–889
Canter Cremers H, Spaink HP, Wijfjes AH, Pees E, Wijffelman CA, Okker RJ, Lugtenberg BJ
(1989) Additional nodulation genes on the Sym plasmid of Rhizobium leguminosarum biovar
viciae. Plant Mol Biol 13:163–174
Cardenas L, Dominguez J, Quinto C, Lopez-Lara IM, Lugtenberg BJ, Spaink HP, Rademaker GJ,
Haverkamp J, Thomas-Oates JE (1995) Isolation, chemical structures and biological activity of
the lipo-chitin oligosaccharide nodulation signals from Rhizobium etli. Plant Mol Biol
29:453–464
Cardenas L, Dominguez J, Santana O, Quinto C (1996) The role of the nodI and nodJ genes in the
transport of Nod metabolites in Rhizobium etli. Gene 173:183–187
22
J.B. Miller and G.E.D. Oldroyd
Cardenas L, Thomas-Oates JE, Nava N, Lopez-Lara IM, Hepler PK, Quinto C (2003) The role of
nod factor substituents in actin cytoskeleton rearrangements in Phaseolus vulgaris. Mol Plant
Microbe Interact 16:326–334
Cardenas L, Martinez A, Sanchez F, Quinto C (2008) Fast, transient and specific intracellular ROS
changes in living root hair cells responding to Nod factors (NFs). Plant J 56:802–813
Cardenas L, Quinto C (2008) Reactive oxygen species (ROS) as early signals in root hair cells
responding to rhizobial nodulation factors. Plant Signal Behav 3:1101–1102
Chabaud M, Genre A, Sieberer BJ, Faccio A, Fournier J, Novero M, Barker DG, Bonfante P (2011)
Arbuscular mycorrhizal hyphopodia and germinated spore exudates trigger Ca2+ spiking in the
legume and nonlegume root epidermis. New Phytol 189:347–355
Chen XC, Feng J, Hou BH, Li FQ, Li Q, Hong GF (2005) Modulating DNA bending affects NodDmediated transcriptional control in Rhizobium leguminosarum. Nucleic Acids Res
33:2540–2548
Cloutler J, Laberge S, Castonguay Y, Antoun H (1996) Characterization and mutational analysis
of nodHPQ genes of Rhizobium sp. strain N33. Mol Plant Microbe Interact 9:720–728
Cook CE, Whichard LP, Turner B, Wall ME, Egley GH (1966) Germination of witchweed (Striga
lutea Lour.): isolation and properties of a potent stimulant. Science 154:1189–1190
Cooper JE (2004) Multiple responses of rhizobia to flavonoids during legume root infection. In:
Callow JA (ed) Advances in botanical research. Academic, London, pp 1–62
Cooper JE (2007) Early interactions between legumes and rhizobia: disclosing complexity in a
molecular dialogue. J Appl Microbiol 103:1355–1365
Corvera A, Prome D, Prome JC, Martinez-Romero E, Romero D (1999) The nolL gene from
Rhizobium etli determines nodulation efficiency by mediating the acetylation of the fucosyl
residue in the nodulation factor. Mol Plant Microbe Interact 12:236–246
D’Haeze W, Holsters M (2002) Nod factor structures, responses, and perception during initiation
of nodule development. Glycobiology 12:79R–105R
Davis EO, Evans IJ, Johnston AW (1988) Identification of nodX, a gene that allows Rhizobium
leguminosarum biovar viciae strain TOM to nodulate Afghanistan peas. Mol Gen Genet
212:531–535
Dazzo FB, Truchet GL, Hollingsworth RI, Hrabak EM, Pankratz HS, Philip-Hollingsworth S,
Salzwedel JL, Chapman K, Appenzeller L, Squartini A et al (1991) Rhizobium lipopolysaccharide modulates infection thread development in white clover root hairs. J Bacteriol
173:5371–5384
Debelle F, Sharma SB (1986) Nucleotide sequence of Rhizobium meliloti RCR2011 genes
involved in host specificity of nodulation. Nucleic Acids Res 14:7453–7472
Debelle F, Plazanet C, Roche P, Pujol C, Savagnac A, Rosenberg C, Prome JC, Denarie J (1996)
The NodA proteins of Rhizobium meliloti and Rhizobium tropici specify the N-acylation of
Nod factors by different fatty acids. Mol Microbiol 22:303–314
Del Papa MF, Pistorio M, Draghi WO, Lozano MJ, Giusti MA, Medina C, van Dillewijn P,
Martinez-Abarca F, Moron Flores B, Ruiz-Sainz JE, Megias M, Puhler A, Niehaus K, Toro N,
Lagares A (2007) Identification and characterization of a nodH ortholog from the alfalfanodulating Or191-like rhizobia. Mol Plant Microbe Interact 20:138–145
Demont N, Debelle F, Aurelle H, Denarie J, Prome JC (1993) Role of the Rhizobium meliloti nodF
and nodE genes in the biosynthesis of lipo-oligosaccharidic nodulation factors. J Biol Chem
268:20134–20142
Denarie J, Debelle F, Truchet G, Prome JC (1993) Rhizobium and legume nodulation: a molecular
dialogue. In: Palacios R, Mora J, Newton W (eds) New horizons in nitrogen fixation. Kluwer,
Dordrecht, pp 19–30
Dennis PG, Miller AJ, Hirsch PR (2010) Are root exudates more important than other sources of
rhizodeposits in structuring rhizosphere bacterial communities? FEMS Microbiol Ecol
72:313–327
Dixon RA, Achnine L, Kota P, Liu CJ, Reddy MS, Wang L (2002) The phenylpropanoid pathway
and plant defence-a genomics perspective. Mol Plant Pathol 3:371–390
The Role of Diffusible Signals in the Establishment of Rhizobial and Mycorrhizal
23
Downie JA (2010) The roles of extracellular proteins, polysaccharides and signals in the
interactions of rhizobia with legume roots. FEMS Microbiol Rev 34:150–170
Ehrhardt DW, Atkinson EM, Long SR (1992) Depolarization of alfalfa root hair membrane
potential by Rhizobium meliloti Nod factors. Science 256:998–1000
Ehrhardt DW, Atkinson EM, Faull KF, Freedberg DI, Sutherlin DP, Armstrong R, Long SR (1995)
In vitro sulfotransferase activity of NodH, a nodulation protein of Rhizobium meliloti required
for host-specific nodulation. J Bacteriol 177:6237–6245
Ehrhardt DW, Wais R, Long SR (1996) Calcium spiking in plant root hairs responding to
Rhizobium nodulation signals. Cell 85:673–681
Felle HH, Kondorosi É, Kondorosi A, Schultze M (1996) Rapid alkalinization in alfalfa root hairs
in response to rhizobial lipochitooligosaccharide signals. Plant J 10:295–301
Felle HH, Kondorosi E, Kondorosi A, Schultze M (2000) How alfalfa root hairs discriminate
between Nod factors and oligochitin elicitors. Plant Physiol 124:1373–1380
Fernandez-Lopez M, D’Haeze W, Mergaert P, Verplancke C, Prome JC, Van Montagu M,
Holsters M (1996) Role of nodl and nodJ in lipo-chitooligosaccharide secretion in
Azorhizobium caulinodans and Escherichia coli. Mol Microbiol 20:993–1000
Fernandez-Lopez M, D’Haeze W, Van Montagu M, Holsters M (1998) Changes in the glycosylation pattern at the reducing end of azorhizobial Nod factors affect nodulation efficiency. FEMS
Microbiol Lett 158:237–242
Firmin JL, Wilson KE, Rossen L, Johnston AWB (1986) Flavonoid activation of nodulation genes
in Rhizobium reversed by other compounds present in plants. Nature 324:90–92
Firmin JL, Wilson KE, Carlson RW, Davies AE, Downie JA (1993) Resistance to nodulation of cv.
Afghanistan peas is overcome by nodX, which mediates an O-acetylation of the Rhizobium
leguminosarum lipo-oligosaccharide nodulation factor. Mol Microbiol 10:351–360
Fisher RF, Tu JK, Long SR (1985) Conserved nodulation genes in Rhizobium meliloti and
Rhizobium trifolii. Appl Environ Microbiol 49:1432–1435
Fisher RF, Long SR (1993) Interactions of NodD at the nod Box: NodD binds to two distinct sites
on the same face of the helix and induces a bend in the DNA. J Mol Biol 233:336–348
Folch-Mallol JL, Marroqui S, Sousa C, Manyani H, Lopez-Lara IM, van der Drift KM, Haverkamp
J, Quinto C, Gil-Serrano A, Thomas-Oates J, Spaink HP, Megias M (1996) Characterization of
Rhizobium tropici CIAT899 nodulation factors: the role of nodH and nodPQ genes in their
sulfation. Mol Plant Microbe Interact 9:151–163
Freiberg C, Fellay R, Bairoch A, Broughton WJ, Rosenthal A, Perret X (1997) Molecular basis of
symbiosis between Rhizobium and legumes. Nature 387:394–401
Geelen D, Mergaert P, Geremia RA, Goormachtig S, Van Montagu M, Holsters M (1993)
Identification of nodSUIJ genes in Nod locus 1 of Azorhizobium caulinodans: evidence that
nodS encodes a methyltransferase involved in Nod factor modification. Mol Microbiol
9:145–154
Geelen D, Leyman B, Mergaert P, Klarskov K, Van Montagu M, Geremia R, Holsters M (1995)
NodS is an S-adenosyl-L-methionine-dependent methyltransferase that methylates chitooligosaccharides deacetylated at the non-reducing end. Mol Microbiol 17:387–397
Geiger O, Spaink HP, Kennedy EP (1991) Isolation of the Rhizobium leguminosarum NodF
nodulation protein: NodF carries a 40 -phosphopantetheine prosthetic group. J Bacteriol 173:
2872–2878
Gibson KE, Kobayashi H, Walker GC (2008) Molecular determinants of a symbiotic chronic
infection. Annu Rev Genet 42:413–441
Giraud E, Moulin L, Vallenet D, Barbe V, Cytryn E, Avarre JC, Jaubert M, Simon D, Cartieaux F,
Prin Y, Bena G, Hannibal L, Fardoux J, Kojadinovic M, Vuillet L, Lajus A, Cruveiller S,
Rouy Z, Mangenot S, Segurens B, Dossat C, Franck WL, Chang WS, Saunders E, Bruce D,
Richardson P, Normand P, Dreyfus B, Pignol D, Stacey G, Emerich D, Vermeglio A, Medigue
C, Sadowsky M (2007) Legumes symbioses: absence of Nod genes in photosynthetic
bradyrhizobia. Science 316:1307–1312
24
J.B. Miller and G.E.D. Oldroyd
Goedhart J, Hink MA, Visser AJ, Bisseling T, Gadella TW Jr (2000) In vivo fluorescence
correlation microscopy (FCM) reveals accumulation and immobilization of Nod factors in
root hair cell walls. Plant J 21:109–119
Gomez-Roldan V, Fermas S, Brewer PB, Puech-Pages V, Dun EA, Pillot J-P, Letisse F, Matusova R,
Danoun S, Portais J-C, Bouwmeester H, Becard G, Beveridge CA, Rameau C, Rochange SF
(2008) Strigolactone inhibition of shoot branching. Nature 455:189–194
Guenoune D, Galili S, Phillips DA, Volpin H, Chet I, Okon Y, Kapulnik Y (2001) The defense
response elicited by the pathogen Rhizoctonia solani is suppressed by colonization of the AMfungus Glomus intraradices. Plant Sci 160:925–932
Gutjahr C, Banba M, Croset V, An K, Miyao A, An G, Hirochika H, Imaizumi-Anraku H,
Paszkowski U (2008) Arbuscular mycorrhiza-specific signaling in rice transcends the common
symbiosis signaling pathway. Plant Cell 20(11):2989–3005
Hamel LP, Beaudoin N (2010) Chitooligosaccharide sensing and downstream signaling: contrasted
outcomes in pathogenic and beneficial plant-microbe interactions. Planta 232:787–806
Hanin M, Jabbouri S, Quesada-Vincens D, Freiberg C, Perret X, Prome JC, Broughton WJ, Fellay R
(1997) Sulphation of Rhizobium sp. NGR234 Nod factors is dependent on noeE, a new hostspecificity gene. Mol Microbiol 24:1119–1129
Harrison MJ (2005) Signaling in the arbuscular mycorrhizal symbiosis. Annu Rev Microbiol
59:19–42
Higgins CF, Hiles ID, Salmond GP, Gill DR, Downie JA, Evans IJ, Holland IB, Gray L, Buckel SD,
Bell AW et al (1986) A family of related ATP-binding subunits coupled to many distinct
biological processes in bacteria. Nature 323:448–450
Honma MA, Ausubel FM (1987) Rhizobium meliloti has three functional copies of the nodD
symbiotic regulatory gene. Proc Natl Acad Sci USA 84:8558–8562
Horvath B, Bachem CW, Schell J, Kondorosi A (1987) Host-specific regulation of nodulation
genes in Rhizobium is mediated by a plant-signal, interacting with the nodD gene product.
EMBO J 6:841–848
Jabbouri S, Fellay R, Talmont F, Kamalaprija P, Burger U, Relic B, Prome JC, Broughton WJ
(1995) Involvement of nodS in N-methylation and nodU in 6-O-carbamoylation of Rhizobium
sp. NGR234 nod factors. J Biol Chem 270:22968–22973
Jabbouri S, Relic B, Hanin M, Kamalaprija P, Burger U, Prome D, Prome JC, Broughton WJ
(1998) nolO and noeI (HsnIII) of Rhizobium sp. NGR234 are involved in 3-O-carbamoylation
and 2-O-methylation of Nod factors. J Biol Chem 273:12047–12055
Jamil M, Charnikhova T, Verstappen F, Bouwmeester H (2010) Carotenoid inhibitors reduce
strigolactone production and Striga hermonthica infection in rice. Arch Biochem Biophys
504:123–131
John M, Rohrig H, Schmidt J, Wieneke U, Schell J (1993) Rhizobium NodB protein involved in nodulation
signal synthesis is a chitooligosaccharide deacetylase. Proc Natl Acad Sci USA 90:625–629
Jones D, Nguyen C, Finlay R (2009) Carbon flow in the rhizosphere: carbon trading at the soil–root
interface. Plant Soil 321:5–33
Kamst E, van der Drift KM, Thomas-Oates JE, Lugtenberg BJ, Spaink HP (1995) Mass spectrometric analysis of chitin oligosaccharides produced by Rhizobium NodC protein in Escherichia
coli. J Bacteriol 177:6282–6285
Kamst E, Pilling J, Raamsdonk LM, Lugtenberg BJ, Spaink HP (1997) Rhizobium nodulation
protein NodC is an important determinant of chitin oligosaccharide chain length in Nod factor
biosynthesis. J Bacteriol 179:2103–2108
Kistner C, Parniske M (2002) Evolution of signal transduction in intracellular symbiosis. Trends
Plant Sci 7:511–518
Kamst E, Bakkers J, Quaedvlieg NE, Pilling J, Kijne JW, Lugtenberg BJ, Spaink HP (1999) Chitin
oligosaccharide synthesis by rhizobia and zebrafish embryos starts by glycosyl transfer to O4
of the reducing-terminal residue. Biochemistry 38:4045–4052
Kobayashi H, Naciri-Graven Y, Broughton WJ, Perret X (2004) Flavonoids induce temporal shifts
in gene-expression of nod-box controlled loci in Rhizobium sp. NGR234. Mol Microbiol
51:335–347
The Role of Diffusible Signals in the Establishment of Rhizobial and Mycorrhizal
25
Koltai H, LekKala SP, Bhattacharya C, Mayzlish-Gati E, Resnick N, Wininger S, Dor E,
Yoneyama K, Hershenhorn J, Joel DM, Kapulnik Y (2010) A tomato strigolactone-impaired
mutant displays aberrant shoot morphology and plant interactions. J Exp Bot 61:1739–1749
Kosuta S, Chabaud M, Lougnon G, Gough C, Denarie J, Barker DG, Becard G (2003) A diffusible
factor from arbuscular mycorrhizal fungi induces symbiosis-specific MtENOD11 expression in
roots of Medicago truncatula. Plant Physiol 131:952–962
Kosuta S, Hazledine S, Sun J, Miwa H, Morris RJ, Downie JA, Oldroyd GE (2008) Differential
and chaotic calcium signatures in the symbiosis signaling pathway of legumes. Proc Natl Acad
Sci USA 105:9823–9828
Krishnan HB, Pueppke SG (1991) Sequence and analysis of the nodABC region of Rhizobium
fredii USDA257, a nitrogen-fixing symbiont of soybean and other legumes. Mol Plant Microbe
Interact 4:512–520
Laeremans T, Caluwaerts I, Verreth C, Rogel MA, Vanderleyden J, Martinez-Romero E (1996)
Isolation and characterization of Rhizobium tropici Nod factor sulfation genes. Mol Plant
Microbe Interact 9:492–500
Larose G, Chênevert R, Moutoglis P, Gagné S, Piché Y, Vierheilig H (2002) Flavonoid levels in
roots of Medicago sativa are modulated by the developmental stage of the symbiosis and the
root colonizing arbuscular mycorrhizal fungus. J Plant Physiol 159:1329–1339
Le Strange KK, Bender GL, Djordjevic MA, Rolfe BG, Redmond JW (1990) The Rhizobium strain
NGR234 nodD1 gene product responds to Activation by the simple phenolic compounds
vanillin and isovanillin present in wheat seedling extracts. Mol Plant Microbe Interact
3:214–220
Lerouge P, Roche P, Faucher C, Maillet F, Truchet G, Prome JC, Denarie J (1990) Symbiotic hostspecificity of Rhizobium meliloti is determined by a sulphated and acylated glucosamine
oligosaccharide signal. Nature 344:781–784
Li F, Hou B, Chen L, Yao Z, Hong G (2008) In vitro observation of the molecular interaction
between NodD and its inducer naringenin as monitored by fluorescence resonance energy
transfer. Acta Biochim Biophys Sin (Shanghai) 40:783–789
Lopez-Lara IM, Blok-Tip L, Quinto C, Garcia ML, Stacey G, Bloemberg GV, Lamers GE,
Lugtenberg BJ, Thomas-Oates JE, Spaink HP (1996) NodZ of Bradyrhizobium extends the
nodulation host range of Rhizobium by adding a fucosyl residue to nodulation signals. Mol
Microbiol 21:397–408
Lopez-Lara IM, Kafetzopoulos D, Spaink HP, Thomas-Oates JE (2001) Rhizobial NodL O-acetyl
transferase and NodS N-methyl transferase functionally interfere in production of modified
Nod factors. J Bacteriol 183:3408–3416
Lopez-Raez JA, Charnikhova T, Gomez-Roldan V, Matusova R, Kohlen W, De Vos R,
Verstappen F, Puech-Pages V, Becard G, Mulder P, Bouwmeester H (2008) Tomato
strigolactones are derived from carotenoids and their biosynthesis is promoted by phosphate
starvation. New Phytol 178:863–874
Lorquin J, Lortet G, Ferro M, Mear N, Dreyfus B, Prome JC, Boivin C (1997a) Nod factors from
Sinorhizobium saheli and S. teranga Sesbania-nodulating strains are both arabinosylated and
fucosylated, a structural feature specific to Sesbania rostrata symbionts. Mol Plant Microbe
Interact 10:879–890
Lorquin J, Lortet G, Ferro M, Mear N, Prome JC, Boivin C (1997b) Sinorhizobium teranga bv.
acaciae ORS1073 and Rhizobium sp. strain ORS1001, two distantly related Acacia-nodulating
strains, produce similar Nod factors that are O carbamoylated, N methylated, and mainly
sulfated. J Bacteriol 179:3079–3083
Mabood F, Souleimanov A, Khan W, Smith DL (2006) Jasmonates induce Nod factor production
by Bradyrhizobium japonicum. Plant Physiol Biochem 44:759–765
Madinabeitia N, Bellogin RA, Buendia-Claveria AM, Camacho M, Cubo T, Espuny MR,
Gil-Serrano AM, Lyra MC, Moussaid A, Ollero FJ, Soria-Diaz ME, Vinardell JM, Zeng J,
Ruiz-Sainz JE (2002) Sinorhizobium fredii HH103 has a truncated nolO gene due to a 1
frameshift mutation that is conserved among other geographically distant S. fredii strains. Mol
Plant Microbe Interact 15:150–159
26
J.B. Miller and G.E.D. Oldroyd
Madsen EB, Madsen LH, Radutoiu S, Olbryt M, Rakwalska M, Szczyglowski K, Sato S, Kaneko T,
Tabata S, Sandal N, Stougaard J (2003) A receptor kinase gene of the LysM type is involved in
legume perception of rhizobial signals. Nature 425:637–640
Maillet F, Poinsot V, Andre O, Puech-Pages V, Haouy A, Gueunier M, Cromer L, Giraudet D,
Formey D, Niebel A, Martinez EA, Driguez H, Becard G, Denarie J (2011) Fungal lipochitooligosaccharide symbiotic signals in arbuscular mycorrhiza. Nature 469:58–63
Marvel DJ, Kuldau G, Hirsch A, Richards E, Torrey JG, Ausubel FM (1985) Conservation of
nodulation genes between Rhizobium meliloti and a slow-growing Rhizobium strain that
nodulates a nonlegume host. Proc Natl Acad Sci USA 82:5841–5845
Matusova R, Rani K, Verstappen FW, Franssen MC, Beale MH, Bouwmeester HJ (2005) The
strigolactone germination stimulants of the plant-parasitic Striga and Orobanche spp. are
derived from the carotenoid pathway. Plant Physiol 139:920–934
McIver J, Djordjevic MA, Weinman JJ, Bender GL, Rolfe BG (1989) Extension of host range of
Rhizobium leguminosarum bv. trifolii caused by point mutations in nodD that result in
alterations in regulatory function and recognition of inducer molecules. Mol Plant Microbe
Interact 2:97–106
Mergaert P, D’Haeze W, Geelen D, Prome D, Van Montagu M, Geremia R, Prome JC, Holsters M
(1995) Biosynthesis of Azorhizobium caulinodans Nod factors. Study of the activity of the
NodABCS proteins by expression of the genes in Escherichia coli. J Biol Chem
270:29217–29223
Mergaert P, D’Haeze W, Fernandez-Lopez M, Geelen D, Goethals K, Prome JC, Van Montagu M,
Holsters M (1996) Fucosylation and arabinosylation of Nod factors in Azorhizobium
caulinodans: involvement of nolK, nodZ as well as noeC and/or downstream genes. Mol
Microbiol 21:409–419
Mergaert P, Ferro M, D’Haeze W, van Montagu M, Holsters M, Prome JC (1997) Nod factors of
Azorhizobium caulinodans strain ORS571 can be glycosylated with an arabinosyl group, a
fucosyl group, or both. Mol Plant Microbe Interact 10:683–687
Mitra RM, Shaw SL, Long SR (2004) Six nonnodulating plant mutants defective for Nod factorinduced transcriptional changes associated with the legume-rhizobia symbiosis. Proc Natl
Acad Sci USA 101:10217–10222
Morón B, Soria-Dı́az ME, Ault J, Verroios G, Noreen S, Rodrı́guez-Navarro DN, Gil-Serrano A,
Thomas-Oates J, Megı́as M, Sousa C (2005) Low pH changes the profile of nodulation factors
produced by Rhizobium tropici CIAT899. Chem Biol 12:1029–1040
Moscatiello R, Squartini A, Mariani P, Navazio L (2010) Flavonoid-induced calcium signalling in
Rhizobium leguminosarum bv. viciae. New Phytol 188:814–823
Mulligan JT, Long SR (1985) Induction of Rhizobium meliloti nodC expression by plant exudate
requires nodD. Proc Natl Acad Sci USA 82:6609–6613
Nagahashi G, Douds DD (1999) Rapid and sensitive bioassay to study signals between root
exudates and arbuscular mycorrhizal fungi. Biotechnology Techniques 13:893–897
Navazio L, Moscatiello R, Genre A, Novero M, Baldan B, Bonfante P, Mariani P (2007) A
diffusible signal from arbuscular mycorrhizal fungi elicits a transient cytosolic calcium elevation in host plant cells. Plant Physiol 144:673–681
Ogawa J, Long SR (1995) The Rhizobium meliloti groELc locus is required for regulation of early
nod genes by the transcription activator NodD. Genes Dev 9:714–729
Olah B, Briere C, Becard G, Denarie J, Gough C (2005) Nod factors and a diffusible factor from
arbuscular mycorrhizal fungi stimulate lateral root formation in Medicago truncatula via the
DMI1/DMI2 signalling pathway. Plant J 44:195–207
Oldroyd GE, Engstrom EM, Long SR (2001a) Ethylene inhibits the Nod factor signal transduction
pathway of Medicago truncatula. Plant Cell 13:1835–1849
Oldroyd GE, Downie JA (2004) Calcium, kinases and nodulation signalling in legumes. Nat Rev
Mol Cell Biol 5:566–576
Oldroyd GE, Downie JA (2006) Nuclear calcium changes at the core of symbiosis signalling. Curr
Opin Plant Biol 9:351–357
The Role of Diffusible Signals in the Establishment of Rhizobial and Mycorrhizal
27
Oldroyd GE, Downie JA (2008) Coordinating nodule morphogenesis with rhizobial infection in
legumes. Annu Rev Plant Biol 59:519–546
Oldroyd GED, Mitra RM, Wais RJ, Long SR (2001b) Evidence for structurally specific negative
feedback in the Nod factor signal transduction pathway. Plant J 28:191–199
Olsthoorn MM, Lopez-Lara IM, Petersen BO, Bock K, Haverkamp J, Spaink HP, Thomas-Oates
JE (1998) Novel branched nod factor structure results from alpha-(1–>3) fucosyl transferase
activity: the major lipo-chitin oligosaccharides from Mesorhizobium loti strain NZP2213 bear
an alpha-(1–>3) fucosyl substituent on a nonterminal backbone residue. Biochemistry
37:9024–9032
Olsthoorn MM, Stokvis E, Haverkamp J, Spaink HP, Thomas-Oates JE (2000) Growth temperature regulation of host-specific modifications of rhizobial lipo-chitin oligosaccharides: the
function of nodX is temperature regulated. Mol Plant Microbe Interact 13:808–820
Op den Camp R, Streng A, De Mita S, Cao Q, Polone E, Liu W, Ammiraju JSS, Kudrna D,
Wing R, Untergasser A, Bisseling T, Geurts R (2011) LysM-type mycorrhizal receptor
recruited for Rhizobium symbiosis in nonlegume parasponia. Science 331:909–912
Ovtsyna AO, Geurts R, Bisseling T, Lugtenberg BJ, Tikhonovich I, Spaink HP (1998) Restriction
of host range by the sym2 allele of Afghan pea is nonspecific for the type of modification at the
reducing terminus of nodulation signals. Mol Plant Microbe Interact 11:418–422
Ovtsyna AO, Rademaker GJ, Esser E, Weinman J, Rolfe BG, Tikhonovich IA, Lugtenberg BJ,
Thomas-Oates JE, Spaink HP (1999) Comparison of characteristics of the nodX genes from
various Rhizobium leguminosarum strains. Mol Plant Microbe Interact 12:252–258
Pacios Bras C, Jorda MA, Wijfjes AH, Harteveld M, Stuurman N, Thomas-Oates JE, Spaink HP
(2000) A Lotus japonicus nodulation system based on heterologous expression of the fucosyl
transferase NodZ and the acetyl transferase NoIL in Rhizobium leguminosarum. Mol Plant
Microbe Interact 13:475–479
Parniske M (2008) Arbuscular mycorrhiza: the mother of plant root endosymbioses. Nat Rev
Microbiol 6:763–775
Peck MC, Fisher RF, Long SR (2006) Diverse flavonoids stimulate NodD1 binding to nod gene
promoters in Sinorhizobium meliloti. J Bacteriol 188:5417–5427
Perret X, Freiberg C, Rosenthal A, Broughton WJ, Fellay R (1999) High-resolution transcriptional
analysis of the symbiotic plasmid of Rhizobium sp. NGR234. Mol Microbiol 32:415–425
Perret X, Staehelin C, Broughton WJ (2000) Molecular basis of symbiotic promiscuity. Microbiol
Mol Biol Rev 64:180–201
Peters NK, Frost JW, Long SR (1986) A plant flavone, luteolin, induces expression of Rhizobium
meliloti nodulation genes. Science 233:977–980
Phillips DA, Joseph CM, Maxwell CA (1992) Trigonelline and Stachydrine released from alfalfa
seeds activate NodD2 protein in Rhizobium meliloti. Plant Physiol 99:1526–1531
Poinsot V, Belanger E, Laberge S, Yang GP, Antoun H, Cloutier J, Treilhou M, Denarie J, Prome
JC, Debelle F (2001) Unusual methyl-branched alpha, beta-unsaturated acyl chain
substitutions in the Nod factors of an arctic rhizobium, Mesorhizobium sp. strain N33
(Oxytropis arctobia). J Bacteriol 183:3721–3728
Price NP, Relic B, Talmont F, Lewin A, Prome D, Pueppke SG, Maillet F, Denarie J, Prome JC,
Broughton WJ (1992) Broad-host-range Rhizobium species strain NGR234 secretes a family of
carbamoylated, and fucosylated, nodulation signals that are O-acetylated or sulphated. Mol
Microbiol 6:3575–3584
Quesada-Vincens D, Fellay R, Nasim T, Viprey V, Burger U, Prome JC, Broughton WJ, Jabbouri
S (1997) Rhizobium sp. strain NGR234 NodZ protein is a fucosyltransferase. J Bacteriol
179:5087–5093
Quesada-Vincens D, Hanin M, Broughton WJ, Jabbouri S (1998) In vitro sulfotransferase activity
of NoeE, a nodulation protein of Rhizobium sp. NGR234. Mol Plant Microbe Interact
11:592–600
Quinto C, Wijfjes AH, Bloemberg GV, Blok-Tip L, Lopez-Lara IM, Lugtenberg BJ, ThomasOates JE, Spaink HP (1997) Bacterial nodulation protein NodZ is a chitin oligosaccharide
28
J.B. Miller and G.E.D. Oldroyd
fucosyltransferase which can also recognize related substrates of animal origin. Proc Natl Acad
Sci USA 94:4336–4341
Radutoiu S, Madsen LH, Madsen EB, Felle HH, Umehara Y, Gronlund M, Sato S, Nakamura Y,
Tabata S, Sandal N, Stougaard J (2003) Plant recognition of symbiotic bacteria requires two
LysM receptor-like kinases. Nature 425:585–592
Radutoiu S, Madsen LH, Madsen EB, Jurkiewicz A, Fukai E, Quistgaard EM, Albrektsen AS,
James EK, Thirup S, Stougaard J (2007) LysM domains mediate lipochitin-oligosaccharide
recognition and Nfr genes extend the symbiotic host range. EMBO J 26:3923–3935
Recourt K, van Brussel AA, Driessen AJ, Lugtenberg BJ (1989) Accumulation of a nod gene
inducer, the flavonoid naringenin, in the cytoplasmic membrane of Rhizobium leguminosarum
biovar viciae is caused by the pH-dependent hydrophobicity of naringenin. J Bacteriol
171:4370–4377
Redmond JW, Batley M, Djordjevic MA, Innes RW, Kuempel PL, Rolfe BG (1986) Flavones
induce expression of nodulation genes in Rhizobium. Nature 323:632–635
Remans R, Snoeck C, Verreth C, Croonenborghs A, Luyten E, Ndayizeye M, Martinez-Romero E,
Michiels J, Vanderleyden J (2007) Inactivation of the nodH gene in Sinorhizobium sp. BR816
enhances symbiosis with Phaseolus vulgaris L. FEMS Microbiol Lett 266:210–217
Ritsema T, Lugtenberg BJ, Spaink HP (1997) Acyl-acyl carrier protein is a donor of fatty acids in
the NodA-dependent step in biosynthesis of lipochitin oligosaccharides by rhizobia. J Bacteriol
179:4053–4055
Roche P, Debelle F, Maillet F, Lerouge P, Faucher C, Truchet G, Denarie J, Prome JC (1991)
Molecular basis of symbiotic host specificity in Rhizobium meliloti: nodH and nodPQ genes
encode the sulfation of lipo-oligosaccharide signals. Cell 67:1131–1143
Roche P, Maillet F, Plazanet C, Debelle F, Ferro M, Truchet G, Prome JC, Denarie J (1996) The
common nodABC genes of Rhizobium meliloti are host-range determinants. Proc Natl Acad
Sci USA 93:15305–15310
Rodpothong P, Sullivan JT, Songsrirote K, Sumpton D, Cheung KW, Thomas-Oates J, Radutoiu S,
Stougaard J, Ronson CW (2009) Nodulation gene mutants of Mesorhizobium loti R7A-nodZ
and nolL mutants have host-specific phenotypes on Lotus spp. Mol Plant Microbe Interact
22:1546–1554
Rohrig H, Schmidt J, Wieneke U, Kondorosi E, Barlier I, Schell J, John M (1994) Biosynthesis of
lipooligosaccharide nodulation factors: Rhizobium NodA protein is involved in N-acylation of
the chitooligosaccharide backbone. Proc Natl Acad Sci USA 91:3122–3126
Rosas S, Soria R, Correa N, Abdala G (1998) Jasmonic acid stimulates the expression of nod genes
in Rhizobium. Plant Mol Biol 38:1161–1168
Sbrana C, Giovannetti M (2005) Chemotropism in the arbuscular mycorrhizal fungus Glomus
mosseae. Mycorrhiza 15:539–545
Scervino JM, Ponce MA, Erra-Bassells R, Vierheilig H, Ocampo JA, Godeas A (2005a)
Flavonoids exhibit fungal species and genus specific effects on the presymbiotic growth of
Gigaspora and Glomus. Mycol Res 109:789–794
Scervino JM, Ponce MA, Erra-Bassells R, Vierheilig H, Ocampo JA, Godeas A (2005b)
Arbuscular mycorrhizal colonization of tomato by Gigaspora and Glomus species in the
presence of root flavonoids. J Plant Physiol 162:625–633
Schlaman HR, Spaink HP, Okker RJ, Lugtenberg BJ (1989) Subcellular localization of the nodD
gene product in Rhizobium leguminosarum. J Bacteriol 171:4686–4693
Schmidt PE, Broughton WJ, Werner D (1994) Nod Factors of Bradyrhizobium japonicum and
Rhizobium sp. NGR234 induce flavonoid accumulation in soybean root exudate. Mol Plant
Microbe Interact 7:384–390
Schultze M, Staehelin C, Rohrig H, John M, Schmidt J, Kondorosi E, Schell J, Kondorosi A (1995)
In vitro sulfotransferase activity of Rhizobium meliloti NodH protein: lipochitooligosaccharide
nodulation signals are sulfated after synthesis of the core structure. Proc Natl Acad Sci USA
92:2706–2709
The Role of Diffusible Signals in the Establishment of Rhizobial and Mycorrhizal
29
Schumpp O, Deakin WJ (2010) How inefficient rhizobia prolong their existence within nodules.
Trends Plant Sci 15:189–195
Schwedock J, Long SR (1990) ATP sulphurylase activity of the nodP and nodQ gene products of
Rhizobium meliloti. Nature 348:644–647
Schwedock JS, Liu C, Leyh TS, Long SR (1994) Rhizobium meliloti NodP and NodQ form a
multifunctional sulfate-activating complex requiring GTP for activity. J Bacteriol
176:7055–7064
Shaw LJ, Morris P, Hooker JE (2006) Perception and modification of plant flavonoid signals by
rhizosphere microorganisms. Environ Microbiol 8:1867–1880
Shaw SL, Long SR (2003) Nod factor elicits two separable calcium responses in Medicago
truncatula root hair cells. Plant Physiol 131:976–984
Smit P, Limpens E, Geurts R, Fedorova E, Dolgikh E, Gough C, Bisseling T (2007) Medicago
LYK3, an entry receptor in rhizobial nodulation factor signaling. Plant Physiol 145:183–191
Sorefan K, Booker J, Haurogne K, Goussot M, Bainbridge K, Foo E, Chatfield S, Ward S,
Beveridge C, Rameau C, Leyser O (2003) MAX4 and RMS1 are orthologous dioxygenaselike genes that regulate shoot branching in Arabidopsis and pea. Genes Dev 17:1469–1474
Soto MJ, Fernández-Aparicio M, Castellanos-Morales V, Garcı́a-Garrido JM, Ocampo JA,
Delgado MJ, Vierheilig H (2010) First indications for the involvement of strigolactones on
nodule formation in alfalfa (Medicago sativa). Soil Biol Biochem 42:383–385
Spaink HP, Wijffelman CA, Pees E, Okker RJH, Lugtenberg BJJ (1987) Rhizobium nodulation
gene nodD as a determinant of host specificity. Nature 328:337–340
Spaink HP, Sheeley DM, van Brussel AA, Glushka J, York WS, Tak T, Geiger O, Kennedy EP,
Reinhold VN, Lugtenberg BJ (1991) A novel highly unsaturated fatty acid moiety of lipooligosaccharide signals determines host specificity of Rhizobium. Nature 354:125–130
Spaink HP, Wijfjes AH, van der Drift KM, Haverkamp J, Thomas-Oates JE, Lugtenberg BJ (1994)
Structural identification of metabolites produced by the NodB and NodC proteins of Rhizobium
leguminosarum. Mol Microbiol 13:821–831
Spaink HP, Wijfjes AH, Lugtenberg BJ (1995) Rhizobium NodI and NodJ proteins play a role in
the efficiency of secretion of lipochitin oligosaccharides. J Bacteriol 177:6276–6281
Stacey G, Luka S, Sanjuan J, Banfalvi Z, Nieuwkoop AJ, Chun JY, Forsberg LS, Carlson R (1994)
nodZ, a unique host-specific nodulation gene, is involved in the fucosylation of the lipooligosaccharide nodulation signal of Bradyrhizobium japonicum. J Bacteriol 176:620–633
Subramanian S, Stacey G, Yu O (2006) Endogenous isoflavones are essential for the establishment
of symbiosis between soybean and Bradyrhizobium japonicum. Plant J 48:261–273
Subramanian S, Stacey G, Yu O (2007) Distinct, crucial roles of flavonoids during legume
nodulation. Trends Plant Sci 12:282–285
Tamasloukht M, Sejalon-Delmas N, Kluever A, Jauneau A, Roux C, Becard G, Franken P (2003)
Root factors induce mitochondrial-related gene expression and fungal respiration during the
developmental switch from asymbiosis to presymbiosis in the arbuscular mycorrhizal fungus
Gigaspora rosea. Plant Physiol 131:1468–1478
Truchet G, Roche P, Lerouge P, Vasse J, Camut S, de Billy F, Prome J-C, Denarie J (1991)
Sulphated lipo-oligosaccharide signals of Rhizobium meliloti elicit root nodule organogenesis
in alfalfa. Nature 351:670–673
Tsai SM, Phillips DA (1991) Flavonoids released naturally from alfalfa promote development of
symbiotic glomus spores in vitro. Appl Environ Microbiol 57:1485–1488
Umehara M, Hanada A, Yoshida S, Akiyama K, Arite T, Takeda-Kamiya N, Magome H,
Kamiya Y, Shirasu K, Yoneyama K, Kyozuka J, Yamaguchi S (2008) Inhibition of shoot
branching by new terpenoid plant hormones. Nature 455:195–200
van Brussel AAN, Bakhuizen R, van Spronsen PC, Spaink HP, Tak T, Lugtenberg BJJ, Kijne JW
(1992) Induction of pre-infection thread structures in the leguminous host plant by mitogenic
lipo-oligosaccharides of Rhizobium. Science 257:70–72
van der Drift KM, Spaink HP, Bloemberg GV, van Brussel AA, Lugtenberg BJ, Haverkamp J,
Thomas-Oates JE (1996) Rhizobium leguminosarum bv. trifolii produces lipo-chitin
30
J.B. Miller and G.E.D. Oldroyd
oligosaccharides with nodE-dependent highly unsaturated fatty acyl moieties. An electrospray
ionization and collision-induced dissociation tandem mass spectrometric study. J Biol Chem
271:22563–22569
Vierheilig H, Bago B, Albrecht C, Poulin MJ, Piché Y (1998) Flavonoids and arbuscular
mycorrhizal fungi. In: Manthey J, Buslig B (eds) Flavonoids in the living system. Plenum,
New York, pp 9–33
Vogel JT, Walter MH, Giavalisco P, Lytovchenko A, Kohlen W, Charnikhova T, Simkin AJ,
Goulet C, Strack D, Bouwmeester HJ, Fernie AR, Klee HJ (2010) SlCCD7 controls
strigolactone biosynthesis, shoot branching and mycorrhiza-induced apocarotenoid formation
in tomato. Plant J 61:300–311
Wais RJ, Keating DH, Long SR (2002) Structure-function analysis of nod factor-induced root hair
calcium spiking in Rhizobium-legume symbiosis. Plant Physiol 129:211–224
Walker SA, Downie JA (2000) Entry of Rhizobium leguminosarum bv.viciae into root hairs
requires minimal Nod factor specificity, but subsequent infection thread growth requires
nodO or nodE. Mol Plant Microbe Interact 13:754–762
Walker SA, Viprey V, Downie JA (2000) Dissection of nodulation signaling using pea mutants
defective for calcium spiking induced by nod factors and chitin oligomers. Proc Natl Acad Sci
USA 97:13413–13418
Wasson AP, Pellerone FI, Mathesius U (2006) Silencing the flavonoid pathway in Medicago
truncatula inhibits root nodule formation and prevents auxin transport regulation by rhizobia.
Plant Cell 18:1617–1629
Westwood JH (2000) Characterization of the Orobanche–Arabidopsis system for studying
parasite–host interactions. Weed Sci 48:742–748
Winkel-Shirley B (2001) Flavonoid biosynthesis. A colorful model for genetics, biochemistry, cell
biology, and biotechnology. Plant Physiol 126:485–493
Yasuda N, Sugimoto Y, Kato M, Inanaga S, Yoneyama K (2003) (+)-Strigol, a witchweed seed
germination stimulant, from Menispermum dauricum root culture. Phytochemistry
62:1115–1119
Yeh KC, Peck MC, Long SR (2002) Luteolin and GroESL modulate in vitro activity of NodD.
J Bacteriol 184:525–530
Yoneyama K, Xie X, Sekimoto H, Takeuchi Y, Ogasawara S, Akiyama K, Hayashi H (2008)
Strigolactones, host recognition signals for root parasitic plants and arbuscular mycorrhizal
fungi, from Fabaceae plants. New Phytol 179:484–494
Yoneyama K, Xie X, Takeuchi Y (2009) Strigolactones: structures and biological activities. Pest
Manag Sci 65:467–470
Yuen JP, Cassini ST, Toledo De Oliveira T, Nagem TJ, Stacey G (1995) Xanthone induction of
nod gene expression in Bradyrhizobium japonicum. Symbiosis 19:131–140
Zaat SA, Schripsema J, Wijffelman CA, van Brussel AA, Lugtenberg BJ (1989) Analysis of the
major inducers of the Rhizobium nodA promoter from Vicia sativa root exudate and their
activity with different nodD genes. Plant Mol Biol 13:175–188
Zhang J, Subramanian S, Stacey G, Yu O (2009) Flavones and flavonols play distinct critical roles
during nodulation of Medicago truncatula by Sinorhizobium meliloti. Plant J 57:171–183
Zuanazzi JAS, Clergeot PH, Quirion J-C, Husson H-P, Kondorosi A, Ratet P (1998) Production of
Sinorhizobium meliloti nod gene activator and repressor flavonoids from Medicago sativa
roots. Mol Plant Microbe Interact 11:784–794
Infection of Lotus japonicus Roots
by Mesorhizobium loti
Katharina Markmann, Simona Radutoiu, and Jens Stougaard
Abstract Like the two important crop legumes soybean and common bean, the
model legume Lotus japonicus develops determinate root nodules. L. japonicus is
normally infected through root hair infection threads in a process closely
synchronised with the progressing primordial cell divisions and organ development. Recent studies of symbiotic mutants have however led to a remarkable and
unexpected discovery of two alternative intercellular infection modes, crack entry
and infection thread independent single cell infection, in L. japonicus. These results
provide genetic support for the origin of rhizobial infection of legumes through
direct intercellular epidermal invasion and indicate that this ancient infection
process in subsequent evolutionary steps was surpassed by the Nod-factor dependent crack entry and root hair infection thread invasions observed in most extant
legumes.
1 Introduction
Soybean (Glycine max) and common bean (Phaseolus vulgaris) are the two most
important crop legumes for the world’s production of food and feed. Like the model
legume Lotus japonicus, both soybean and bean form determinate root nodules
following inoculation with their respective rhizobial microsymbionts. A determinate nodule is characterised by a transient meristematic activity and cessation of
infection thread growth after the majority of cells in the central zone of the
primordia have been infected. The formation of a determinate root nodule is
therefore a sequential process where clearly defined developmental stages replace
each other and can thus be separated in time. In contrast to determinate nodules,
K. Markmann • S. Radutoiu • J. Stougaard (*)
Department of Molecular Biology, Centre for Carbohydrate Recognition and Signalling (CARB),
Aarhus University, Gustav Wieds Vej 10, 8000 Aarhus C, Denmark
e-mail: [email protected]
S. Perotto and F. Baluska (eds.), Signaling and Communication in Plant Symbiosis,
Signaling and Communication in Plants 11, DOI 10.1007/978-3-642-20966-6_2,
# Springer-Verlag Berlin Heidelberg 2012
31
32
K. Markmann et al.
which tend to be round at maturity, indeterminate nodules of for example pea,
clover and a second model legume Medicago truncatula retain meristematic activity near the nodule tip and therefore develop into elongate structures. Underlying
the tip meristem is the infection zone, where newly produced nodule cells are
infected. Following the early stages founding the nodule primordium, an individual
indeterminate root nodule will therefore represent spatially – but not temporally –
separated stages.
Legume root nodule formation results from two tightly coordinated processes
running in parallel. Development of nodule primordia from dedifferentiated root
cortical cells initiates a meristem, which will later form the nodule. Simultaneously,
a bacterial invasion process targets the primordia. In soybean and bean the outer
cortical cells initiate primordia formation. In L. japonicus initial cell divisions were
detected in cells of both the outer and middle cortex (Szczyglowski et al. 1998; van
Spronsen et al. 2001). Cytoplasmic bridges preparing cells for infection thread
passage were observed associated with primordia in L. japonicus, but not in bean
(van Spronsen et al. 2001). Like soybean and bean, L. japonicus is normally
infected through root hair infection threads, although capacities for crack entry
and intercellular invasion were recently uncovered in studies of mutants (Karas
et al. 2005; Groth et al. 2010; Madsen et al. 2010). The bacterial infection process
starts with rhizobial attachment to plant root hair tips. Subsequent physiological and
morphological responses of root hairs result in their curling, and bacteria are
entrapped in an infection pocket at the curls’ centre (Szczyglowski et al. 1998;
Oldroyd and Downie 2004; Miwa et al. 2006). Starting from the pocket, inwards
growing infection threads form and are colonised by rhizobia. These infection
threads act as rhizobial conduits from which the Mesorhizobium loti symbiont
ultimately will be released and endocytosed into nodule primordia cells. This
infection mode targets pre-developed cells of the emerging nodule, and uninfected,
so-called interstitial cells, are retained in between the infected cells of the central
zone (Imaizumi-Anraku et al. 1997). As a result, infected cells constitute only a
subset of cells in the central zone of determinate L. japonicus, bean and soybean
nodules. In the infected cells rhizobia differentiate into bacteroids that are
surrounded by a peribacteriod membrane derived from the infection thread membrane during endosymbiotic uptake. Together, bacteroids and membrane are
referred to as symbiosomes, organelle-like structures where atmospheric dinitrogen
is reduced to ammonium. Access to this reduced form of nitrogen provides the host
with an ecological advantage over other plants under conditions where soil nitrogen
resources are limiting. The M. loti bacteroids contained in L. japonicus nodules are
not terminally differentiated and maintain their normal bacterial size, genome
content and ability to divide (Van de Velde et al. 2010). This is in contrast to the
indeterminate nodules of M. truncatula, where irreversible differentiation of
bacteriods is thought to be promoted by an abundance of nodule-specific cysteine-rich anti microbial peptides (Van de Velde et al. 2010). L. japonicus nodules are
devoid of these anti microbial peptides, which could be important in maintaining
the continuous infection process in the indeterminate nodules (Van de Velde et al.
2010).
Infection of Lotus japonicus Roots by Mesorhizobium loti
33
2 Recognition Precedes Infection
Several bacterial species belonging to both the a- and the ß-proteobacteria (Moulin
et al. 2001; Broughton 2003) can induce root nodules on different legume species.
The relationship between the approximately 18,000 legume species and their
bacterial microsymbionts is nevertheless selective. L. japonicus can for example
develop fully functional nitrogen fixing nodules with M. loti and ineffective or
partially effective nodules with NGR234, Rhizobium etli and Bradyrhizobium spp.
(Cardenas et al. 1995; Hussain et al. 1999; Banba et al. 2001; Schumpp et al. 2009;
Bek et al. 2010). Traditionally these host/non-host relationships were described in
the so-called cross-inoculation groups composed of legumes and their compatible
rhizobia. However, this catalogue system is inconsistent and far from a perfect
reflection of the host/non-host relationships as elucidated using molecular methods.
Ongoing efforts to fine-tune our understanding of host-microbiont specificity
and the recognition process at the molecular level discovered a two-way signal
exchange as one of the central components of recognition. Rhizobial NodD proteins
mediate host recognition by interacting with specific flavonoids or isoflavonoids
secreted from host roots (Peters et al. 1986; Spaink et al. 1989). Flavonoid-activated
NodD promotes transcription of bacterial nod-genes involved in synthesis and
secretion of lipochitin-oligosaccharides (Nod-factors). These molecules serve as
major bacterial signals detected by the legume host (Mulligan and Long 1985;
Spaink et al. 1991; Truchet et al. 1991). This signalling mechanism was elegantly
demonstrated using two bacteria, R. etli and M. loti, from different cross-inoculation groups nodulating bean and Lotus, respectively. Both strains synthesise Nodfactors with the same structure. Artificially bypassing the flavonoid NodD activation by expression of a constitutive nodD transcriptional activator (FITA) in these
rhizobial strains extended their host range beyond their cross-inoculation group to
encompass both bean and Lotus corniculatus (Cardenas et al. 1995).
3 Nod-Factor Perception and Its Role in Host Determination
In legumes, the lipochitin-oligosaccharide Nod-factors of compatible bacteria initiate a signal cascade leading to nodulin gene activation, root cortical cell dedifferentiation and development of the root nodule (Spaink et al. 1991; Truchet et al.
1991; Hoegslund et al. 2009). The length of the Nod-factor carbohydrate moiety,
the size and degree of saturation of the acyl chain and substitutions of the reducing
and non-reducing glucosamine residues are characteristic for each rhizobial species. It is these structural features that determine whether the Nod-factor is perceived by the plant (Lerouge et al. 1990; Spaink et al. 1991; Truchet et al. 1991).
Mesorhizobium loti strain R7A produces Nod-factors with pentameric GlcNAc
backbones. At the reducing end moiety there is a 4-O-fucose with either an acetyl
group or a proton in position 3 or 4. The nonreducing moiety is N-methylated and
34
K. Markmann et al.
N-acylated (cis-vaccenic acid or stearic acid), and a carbamoyl group is present in
position 3 (Lopez-Lara et al. 1995; Bek et al. 2010). Phenotypic characterisation of
mutants of M. loti strain R7A show an unchanged infection process and nodule
formation in the absence of the N-Methyl and the carbamoyl substitutions. In
contrast, the presence of the N-acetylated fatty acid was found to be crucial for
nodulation. The acetylated fucosyl group was important for effective Nod-factor
signalling and absence of this group led to host dependent nodulation phenotypes
when comparing the four Lotus species L. japonicus, L. filicaulis, L. corniculatus
and L. burttii (Rodpothong et al. 2009).
Perception of Nod-factor in L. japonicus is mediated by receptor kinases
containing LysM modules in their extracellular domains. The two receptor kinases
perceiving the Nod-factor signal, NFR1 and NFR5, are predicted to have a topology
where single pass transmembrane domains anchor LysM containing extracellular
domains and intracellular serine/threonine protein kinases (Madsen et al. 2003;
Radutoiu et al. 2003). Combining this prediction with genetic evidence, a
heteromeric receptor complex composed of NFR1 and NFR5 was proposed to
initiate signal transduction following perception of a correctly decorated Nodfactor (Radutoiu et al. 2003). Biochemical studies have subsequently demonstrated
NFR1 to be a dual specificity kinase capable of autophosphorylation and phosphorylation of NFR5 (Madsen et al. 2011). In contrast, no kinase activity could be
detected for NFR5, which has domain features characteristic for pseudokinases.
Membrane localisation and interaction between NFR1 and NFR5 was shown
in vitro by enrichment in purified membrane fractions and in vivo through bimolecular fluorescence complementation (BiFC) in Nicotiana benthamiana and leek
cells (Madsen et al. 2011).
The structural similarity between Nod-factors and chitin, a carbohydrate
synthesised by different plant pathogens such as fungi, nematodes or insects,
raise the possibility of an evolutionary link between chitin and Nod-factor perception, and thus between symbiotic and pathogenetic interactions. In support of this
hypothesis, the Arabidopsis chitin receptor CERK1, which was shown to be
involved in the perception of fungal pathogens (Miya et al. 2007; Wan et al.
2008), shows high sequence and structural similarity to the NFR1 protein (Radutoiu
et al. 2003; Miya et al. 2007; Wan et al. 2008). Subsequent functional studies of the
kinase domains further supported this close relationship. Exchange of the NFR1
kinase domain with the corresponding CERK domain and substitution of a portion
of the aEF helix in CERK1 with the YAQ amino acid sequence from NFR1
reconstituted symbiotic signalling. Transgenic L. japonicus nfr1 mutant plants
carrying this swap construct were functionally complemented and able to develop
nodules with M. loti (Nakagawa et al. 2011).
A deeper understanding of the host/non-host relationship between rhizobia and
legumes was obtained by transforming the L. japonicus NFR1 and NFR5 genes into
M. truncatula. Using this approach it was shown that NFR1 and NFR5 receptors act
in concert as host determinants, transforming the non-host M. truncatula into a host
able to recognise and be infected by M. loti and an engineered Rhizobium
leguminosarum DZL producing a Nod-factor substituted with an acetylated fucosyl
Infection of Lotus japonicus Roots by Mesorhizobium loti
35
on the reducing moiety (Radutoiu et al. 2007). Recognition of these normally noncompatible bacteria triggers root cell dedifferentiation, redifferentiation and initiation of nodule organogenesis as well as infection thread formation. This extended
NFR1 and NFR5 mediated signal cascade is dependent on both Nod-factor synthesis, as shown with the M. loti nodC mutant, and structure as shown by the longer
infection threads obtained with R. leguminosarum DZL, respectively. In line with
these results it was shown that the L. japonicus NFR1 and NFR5 proteins also
act in concert if introduced into Lotus filicaulis. Transgenic roots of this
species expressing the L. japonicus receptor molecules can be nodulated by the
R. leguminosarum DZL strain, which can nodulate L. japonicus, but is otherwise
unable to nodulate wild type L. filicaulis roots (Radutoiu et al. 2007).
4 The Role of LysM Domains
Our understanding of the mechanisms involved in Nod-factor perception
was further refined through domain swap experiments using receptors from
L. japonicus and L. filicaulis. Initially it was observed that L. filicaulis, in contrast
to L. japonicus, did not develop root nodules after inoculation with the
R. leguminosarum DZL strain (Pacios-Bras 2003). Analysis of transgenic plants
subsequently traced this inability back to variations in the amino acid composition
of the NFR1 or NFR5 receptors between L. japonicus and L. filicaulis (Radutoiu
et al. 2007). Domain swaps combined with substitutions of single amino acids were
then used to show that specific recognition of the DZL Nod-factor relied on the
LysM containing domains of NFR1 and NFR5 and that the LysM2 domain of NFR5
played a major role in discriminating M. loti and R. leguminosarum DZL Nodfactors in L. filicaulis. A leucine adjacent to a putative Nod-factor binding groove
located between the first b-strand and first helix of the L. japonicus NFR5 LysM2
domain, a position filled by a lysine in L. filicaulis, was found to be largely
responsible for the recognition of the Nod-factor synthesised by the DZL strain
(Radutoiu et al. 2007). However, presence of three LysM domains in the NFR5 and
NFR1 receptors (Madsen et al. 2003; Radutoiu et al. 2003; Arrighi et al. 2006),
suggests the involvement of more than one LysM domain in Nod-factor perception.
Two lines of evidence supports this notion: (1) the non-nodulation phenotype
caused by an amino acid substitution in the LysM1 domain of the M. trunctula
homolog of NFR5 called NFP (Arrighi et al. 2006) and (2) the involvement of
LysM1 of the pea SYM37 NFR1-like receptor in distinguishing “European” and
“Middle East” R. leguminosarum bv. viciae strains (Zhukov et al. 2008).
Further insight into the functional role of individual amino acids in Nod-factor
perception was obtained in a domain swap study using the extracellular domains of
NFR1 and NFR5 from a more distantly related species, Lotus pedunculatus. This
Lotus species is normally nodulated by a Bradyrhizobium spp. strain producing a
Nod-factor with an additional carbamoyl group at the non-reducing moiety (Bek et al.
2010). It was found that the combined amino acid differences of the NFR1 and NFR5
36
K. Markmann et al.
extracellular domains of L. japonicus and L. pedunculatus were not influencing the
recognition of the Nod-factor substituted with one or two carbamoyls at the nonreducing end. Considering that a high number of amino acid variations is found
between L. japonicus and L. pedunculatus NFR proteins, this suggests the involvement of only a small fraction of amino acids in deciphering Nod-factor structure.
5 Signal Transduction and Signalling Triggered
by the NFR Receptors
In L. japonicus signal transduction following NFR-mediated Nod-factor perception
is shared with the arbuscular mycorrhizal symbiosis. Common symbiotic signalling
components include at least eight genes encoding the leucine-rich repeat RLK
SYMRK (Stracke et al. 2002), the cation channels CASTOR and POLLUX
(Imaizumi-Anraku et al. 2005), the nuclear pore proteins NUP133 (Kanamori
et al. 2006), NUP85 (Saito et al. 2007) and NENA (Groth et al. 2010), the Ca2+/
calmodulin dependent kinase CCaMK (Tirichine et al. 2006) and the nuclear
protein CYCLOPS (Yano et al. 2008). Analysis of L. japonicus mutants has
shown that the LysM containing Nod-factor receptor(s), SYMRK, CASTOR and
POLLUX, and the nuclear pore proteins are required for the induction of calcium
oscillations, one of the earliest physiological responses detectable in root hairs after
exposure to purified Nod-factor (Miwa et al. 2006; Saito et al. 2007). These calcium
oscillations are thought to be interpreted by the CCaMK encoded Ca2+/calmodulin
dependent protein kinase (Miwa et al. 2006; Tirichine et al. 2006). A function in
cross signalling between infection and organogenesis was proposed for CYCLOPS
(Madsen et al. 2010), a direct interactor of CCaMK (Messinese et al. 2007; Yano
et al. 2008). Downstream of the shared symbiosis pathway, putative transcription
factors encoded by Nin (Schauser et al. 1999), Nsp1 and Nsp2 (Heckmann et al.
2006), and a member of the ERF family (Asamizu et al. 2008; Middleton et al.
2007) are required for regulation of nodule expressed genes and initiation
of nodule organogenesis. One of these transcripion factors, NSP2, has been proposed to also contribute to arbuscular mycorrhizal symbiosis in M. truncatula
(Maillet et al. 2011). This implies the interesting possibility that common symbiotic
signalling could include the level of transcription-factor mediated downstream gene
activation.
Formation of nodule primordia involves dedifferentiation and reactivation of
cortical root cells. A combination of gain and loss of function mutants have shown
that cytokinin signalling through the Lhk1 cytokinin receptor is necessary and
sufficient for the formation of nodule primordia (Murray et al. 2007; Tirichine
et al. 2007). The phenotype of loss of function mutants (hit1) includes a drastic
reduction in the number of primordia initiated, suggesting that cytokinin signalling
is important for activation of the cell cycle in inner cortical foci that form the starting
point of nodule organogenesis (Murray et al. 2007). This phenotype complies with
Infection of Lotus japonicus Roots by Mesorhizobium loti
37
the detection of increased expression of a cytokinin reporter gene in the early phases
of nodulation (Lohar et al. 2004). In the dominant snf2 mutants encoding a gain of
function LHK1 cytokinin receptor, the opposite phenotype is observed. snf2 mutants
develop root nodules spontaneously, i.e. in the absence of any rhizobia or Nodfactor signalling (Tirichine et al. 2007). This demonstrates that cytokinin signalling
can induce the nodule meristem directly, presumably by activation of the cell cycle
in distinct cell foci along the root. Both of these Lhk1 mutants therefore affect the
developmental events in the root cortex, suggesting that cytokinin is a component of
the secondary epidermal-cortical signalling triggered by Nod-factor perception. The
epidermal developments like root hair deformation and infection thread formation
appear to be mainly indirectly affected by cytokinin signalling. In hit1 loss of
function mutants, hyperinfection in the form of an increased number of root hair
infection threads are observed, and these infection threads are restricted in their
progression into the cortex (Murray et al. 2007). This suggests that nodule primordia
are involved in regulating infection thread formation through a negatively acting
mechanism, and are necessary for their controlled progression into the inner cortex.
The disruption of infection thread progression into the cortex in the hit1 loss of
function mutant further indicates a positive role for cytokinin signalling. Cytokinin
is thus a candidate signal for the coordination of epidermal and cortical processes of
infection and primordium formation, respectively.
6 Cell to Cell Recognition Mechanisms Regulate
Bacterial Entry
Early investigations suggested plant lectin binding of bacterial specific extracellular
carbohydrates to be the major determinants of plant-rhizobium host range (Bohlool
and Schmidt 1974). Subsequent experiments demonstrated binding of legume lectins
to rhizobial surface polysaccharides (Bourne et al. 1994), and indicated that lectins
facilitate rhizobial attachment to the root hair tips by binding surface polysaccharides
and thereby increasing the local Nod-factor concentration above the required threshold for nodule initiation (van Rhijn et al. 2001). Studies performed with bacterial
mutants showed that lectin mediated recognition is particularly important for infection thread progression through cortical cell layers (van Rhijn et al. 1998). Rhizobial
host range extension as a result of lectin gene transfer from pea or soybean into
clover, alfalfa or L. corniculatus appears also in most cases to be dependent on the
Nod-factor structure and/or bacterial exopolysaccharides (van Rhijn et al. 1998,
2001). The mechanism and contribution of plant lectins to the infection process is
therefore awaiting further investigation. For a recent review summarising the role of
lectins in plant-microbe interactions see De Hoff et al. (2009).
The precise role of the different rhizobial surface polysaccharides for symbiotic
partner recognition of bacteria and/or for the complete infection of the plant root
tissues is on the other hand, well documented through genetic analysis of rhizobia.
38
K. Markmann et al.
cgs mutants of M. loti R7A or strain Ayac 1 BII that are deficient in the synthesis of
cyclic ß-glucan induced the formation of empty nodules on L. japonicus (Kelly,
personal communication; D’Antuono et al. 2005, 2008). Infection thread development was impaired indicating a role for cyclic ß-glucan during infection thread
formation or progression. The lipopolysaccharide (LPS) deficient lps mutants of
M. loti tested on L. japonicus so far were only marginally affected in establishing
symbiosis, and the phenotypes observed are mainly associated with strain competitiveness (Kelly, personal communication; D’Antuono et al. 2008). Further studies
are required in order to determine the influence of LPS on symbiosis in
L. japonicus, which in light of the recent report of nitric oxide (NO) release in
response to purified LPS may be complex (Murakami et al. 2011). Exopolysaccharides (EPS) appear to be more important for infection of L. japonicus. Nodulation of L. japonicus by eps mutant strains defective in genes that are involved
at mid-late stages of EPS biosynthesis (exoU, exoO, exoK and mlr5265) was
severely affected. These mutant bacteria induced the formation of small white
bumps devoid of bacteria. However, an EPS mutant strain (exoB) disrupted in
a very early stage of EPS biosynthesis forms nitrogen-fixing nodules indistinguishable from those induced by wild-type M. loti. Visualisation of the exoU mutant
bacteria during the infection process revealed that the EPS-deficient strain was
disrupted in its ability to induce normal infection thread formation. The host plant
may therefore be able to decipher the EPS structure presented by the invading
bacteria most likely through a specialised EPS receptor (Kelly, personal communication). A specific role for bacterial EPS is further supported by an earlier study
showing that several eps mutant strains of M. loti strain PN184 lose compatibility
with the host Leucaena leucocephala, while retaining full compatibility with
a second host, L. pedunculatus (Hotter and Scott 1991).
Strains of M. loti possess either type III or type IV (e.g. MAFF 303099 and R7A,
respectively) secretion systems that in several cases have been shown to deliver
effector proteins into cells of eukaryotic hosts and to be required for virulence of
many Gram-negative bacterial pathogens. In the M. loti strain R7A, expression of
genes encoding components of the type IV secretion system is specifically
regulated in a symbiosis-dependent manner and linked to the presence of the
host (Hubber et al. 2004, 2007). Inactivation of the type III secretion system in
the M. loti strain MAFF affects symbiosis neutrally, positively or negatively
dependent on the Lotus species tested (Sanchez et al. 2009; Okazaki et al. 2010).
L. japonicus was unaffected, while L. filicaulis and L. coniculatus subsp. frondosus
formed fewer root nodules in the absence of the secretion system. In contrast, there
was a strong negative effect on infection of L. halophilus, which appears capable of
detecting a small secreted bacterial effector protein and halting symbiosis development in response (Okazaki et al. 2010). Mutation of the gene encoding this protein
reverts the phenotype to fully functional root nodules. Drastic phenotypic changes
resulting from mutation of the type III and IV secretion system were found on the
alternative host L. leucocephala (Hubber et al. 2004, 2007; Okazaki et al. 2010).
Both the R7A and MAFF wild type strains of M. loti induced tumour-like nodules
on this host. Inactivation of both the type III and IV protein effector secretion
Infection of Lotus japonicus Roots by Mesorhizobium loti
39
systems in R7A and MAFF lead to the development of fully infected and functional
root nodules. Recognition of effector proteins resulting in either suppression of
defence responses or induction of the defence response following detection of
secreted proteins as pathogen effectors could explain these opposite responses
(Hubber et al. 2004, 2007). Interestingly, a gene encoding a Toll-interleukin
leucine-rich repeat receptor (TIR-NBS-LRR) suggested to recognise a rhizobial
protein effector was identified in soybean (Yang et al. 2010).
7 Invasion Through Root Hair Infection Threads
In L. japonicus the normal infection occurs through infection threads initiated in
elongating root hairs of the susceptible region located just behind the root tip. Upon
formation of infection pockets, the root hair cell wall dissolves and an infection
thread is initiated by invagination and subsequent polar extension of the plasma
membrane, which is accompanied by the deposition of new cell wall material (see
Gage (2004) for a review). Inward growing infection threads progress through the
root hair and are propagated through the cell layers by a reiterated cell autonomous
mechanism. Such infection threads are only formed in the presence of rhizobial
bacteria. Nod-factor alone elicits the earliest detectable responses such as membrane depolarisation, ion fluxes across the membrane, Ca2+ spiking and cellular
alteration in actin and microtubule organisation, but not infection threads
(Weerasinghe et al. 2003, 2005; Vassileva et al. 2005; Miwa et al. 2006). Interestingly, a class of mutants characterised by a lack of or reduced infection thread
progression have been identified (Schauser et al. 1998; Lombardo et al. 2006; Yano
et al. 2006; Murray et al. 2007). In these mutant lines the infection threads were
typically arrested either within root hairs/epidermis or within the first cortical cell
layers. Furthermore, mutant lines were also identified where infection thread
formation was not followed by the release of bacteria into cells of the nodule
primordium (Imaizumi-Anraku et al. 1997).
Mutant lines known to affect infection thread formation include nin (Schauser
et al. 1999), nsp1 and nsp2 (Heckmann et al. 2006), cerberus (Yano et al. 2009),
nap1 and pir1 (Yokota et al. 2009), alb1-1 (Imaizumi-Anraku et al. 2000), crinkle
(Tansengco et al. 2003), as well as itd1, itd3 and itd4 (Lombardo et al. 2006).
Around half of these have been characterised. A functional Nin gene is required for
infection thread initiation and for restricting root hair deformation, as well as
restricting the size of the infection zone. The domain structure of the NIN protein
suggests a function as transcriptional regulator, and the mutant phenotype
demonstrates an essential role for the protein in both infection thread formation
and organ initiation (Schauser et al. 1999). The phenotype and the transcripts
affected in this mutant background also suggest the involvement of NIN in both
positive and negative regulation and coordination of infection and organogenesis
(Schauser et al. 1999; Hoegslund et al. 2009). Like NIN, NSP1 and NSP2 appear to
be involved in activating the gene expression required for infection thread
40
K. Markmann et al.
formation. The L. japonicus gene set controlled by these regulators has so far not
been identified, but detailed analysis of the putative M. truncatula orthologs of
NSP1 and NSP2 suggests that they act as a transcription factor complex activating
several downstream genes, including NIN and the symbiosis-induced gene
ENOD11 (Hirsch et al. 2009). L. japonicus plants carrying a mutant nap1 or pir1
allele had a significantly diminished ability to capture bacteria within infection
pockets and initiating infection threads (Yokota et al. 2009). The rare infection
threads that formed disintegrated, and infection threads extending to the base of the
root hair cells were only occasionally observed. Lack of infection threads in the root
cortex further suggested that NAP1 and PIR1 were essential for the progression of
the infection process. Further characterisation showed that the Nap1 and Pir1 genes
are essential for establishing the actin organisation in root hairs (Yokota et al.
2009). Molecular analysis revealed the putative Arabidopsis orthologs of these
proteins, NAP1 and PIR1, as likely components of the SCAR/WAVE complex
that activates the ARP2/3 complex, which binds pre-existing actin polymers and
nucleates new actin filaments (Brembu et al. 2004; Deeks et al. 2004; Li et al.
2004). The role of the Cerberus encoded ubiquitin-E3-ligase for infection thread
formation is less clear, but its requirement suggests that there is a need for clearing
the root hair of particular proteins in order to secure infection thread progression
(Yano et al. 2009). A membrane raft associated remorin belonging to a subgroup
found so far only in plants of the Rosid I clade, which all nodulators are part
of, is upregulated during nodulation (Lefebvre et al. 2010). In M. truncatula this
remorin interacts with symbiotic receptors and localises to infection thread- and
symbiosome-membranes (Lefebvre et al. 2010). Functionally the remorin was
linked to a role in bacterial release and endocytosis, and symbiosis was impaired
in remorin mutant plants. So far remorin mutants have not been isolated in
L. japonicus, but it will be interesting to investigate the role of this putative
membrane anchor for the NFR receptors in determinate nodulation. Similar roles
have been suggested for flotillins (Haney and Long 2010) and Rab proteins
(Limpens et al. 2009) within indeterminate nodules of M. truncatula. Different
Rab protein encoding mRNAs have been shown to be upregulated in L. japonicus
nodules (Borg et al. 1997), and analysis of these, as well as of L. japonicus
homologs of remorins and flotillins may further contribute to a better understanding
of infection during determinate root nodule formation.
8 Parallel Signalling Pathways Regulate Root Hair
Infection and Nodule Development
Bacterial infection of legume root nodule primordia is tightly controlled and closely
synchronised with the progressing primordial cell divisions and organ development. This coordination has long prevented the separation of the molecular
mechanisms underlying these two developmental processes and has limited the
Infection of Lotus japonicus Roots by Mesorhizobium loti
41
identification of plant genes controlling the bacterial infection process, but not
nodule organogenesis. As a consequence the genetics of infection thread development is fairly undescribed and among the genes defining this pathway, only Nap1,
Pir1 and Cerberus have so far been characterised at the molecular level (Yano et al.
2009; Yokota et al. 2009). Most of the Nod-factor signal transduction pathway
mutants mentioned above suffer from simultaneous absence or severe impairment
of both organ formation and infection thread development. This has made the
dissection of direct and indirect mutational impact on the two processes very
difficult. However, combining gain of function mutations with loss of function
mutations in trangenes or double, triple and quadruple mutants opened new
possibilities for assessing the role of the genes inactivated by loss of function
mutations (Hayashi et al. 2010; Madsen et al. 2010). The autoactive versions of
CCaMK and LHK1 encoded by the snf1 or snf2 alleles, respectively, were key to
this approach. Both snf mutants form spontaneous nodules independent of bacterial
presence or infection (Tirichine et al. 2006, 2007). Exploiting the ability of these
gain of function alleles to activate the developmental processes from downstream
positions defined the backbones of two parallel pathways facilitating (1) infection
thread formation and (2) root nodule organogenesis (Hayashi et al. 2010; Madsen
et al. 2010). Furthermore, the approach revealed cross-signalling functions between
these two pathways for some of the examined genes. It was shown that the LRR
receptor kinase SYMRK, the nucleoporins NUP133 and NUP85 and the cation
channels CASTOR and POLLUX, all involved in signal transduction down-stream
of Nod-factor perception, were dispensable for root hair infection thread development and invasion of nodule primordia in a snf1 genetic background. Since inactivation of the LRR receptor kinase SYMRK, the nucleoporins and the cation
channels results in absence of calcium spiking (Miwa et al. 2006), these results
further indicate that calcium spiking, apart from activation of CCaMK, is dispensable for infection thread formation (Madsen et al. 2010).
On the other hand, the NFR1 and NFR5 Nod-factor receptors were required for
root hair infection thread initiation, and the NAP1 and PIR1 proteins mediating
actin rearrangement together with the CERBERUS ubiquitin E3 ligase were
required for infection thread progression (Madsen et al. 2010). These observations
support an early branching of the Nod-factor signal transduction pathway. The NIN,
NSP1 and NSP2 transcriptional regulators were required for both infection thread
formation and organogenesis, supporting a simultaneous or sequential role of these
proteins in both of these processes. The absence of infection in cyclops snf1 double
mutants (Madsen et al. 2010) along with the reported protein-protein interaction
between CCaMK and CYCLOPS (Messinese et al. 2007; Yano et al. 2008),
indicated a role for CYCLOPS in cross signalling between the organogenic and
infection pathways (Madsen et al. 2010).
Nod-factor induced root hair deformation, Ca2+ influx and Ca2+ spiking preceding infection thread formation all require the initial NFR1 and NFR5 mediated
perception event (Radutoiu et al. 2003; Miwa et al. 2006). The relation between
these phenomena is however not yet fully clarified. Ca2+ spiking following application of Nod-factors is not required for root hair deformation (Miwa et al. 2006)
42
K. Markmann et al.
and the presumed Ca2+ dependent activation of CCaMK as mimicked by the snf1
mutation seems insufficient for root hair infection thread formation (Madsen et al.
2010). Separation of root hair deformation and influx was also implied by the
observation that three orders of magnitude lower Nod-factor concentration was
sufficient to induce root hair deformation compared with that required for the Ca2+
influx (Radutoiu et al. 2003; Miwa et al. 2006). This could suggest that the NFR
receptor dependent Ca2+ influx observed in root hairs at high Nod-factor concentration may be a prerequisite for root hair infection thread formation. So far these
observations are consistent with the Ca2+ responses occurring in parallel with
the NAP1 and PIR1 induced cytoskeletal changes, although the possibility of
a low level or localised change in Ca2+ cannot be excluded.
9 Alternative Infection Mechanisms: Crack Entry
Microscopical surveys of infection in legume species representing different clades
of the legume family have revealed the existence of a number of alternative
infection modes leading to nitrogen fixing root nodules (recently reviewed in
Held et al. (2010)). However, with only a few exceptions like Sesbania rostrata
(Goormachtig et al. 2004), generally only one infection mode is found in an
individual species. L. japonicus is, for example, infected via intracellular root
hair infection threads (Szczyglowski et al. 1998; Schauser et al. 1999; Lombardo
et al. 2006), while peanut is infected via an intercellular crack entry process (Fabra
et al. 2010). One of the surprising outcomes of the experiments combining gain of
function and loss of function mutations was the discovery of two additional,
although less effective, intercellular infection modes in L. japonicus (Madsen
et al. 2010). Loss-of-function alleles of particular genes also indicated the presence
of a crack entry based infection mode (Karas et al. 2005; Groth et al. 2010), and
mutational abrogation of both root hair and cortical infection thread formation
revealed a further mode of infection of presumably more ancient evolutionary
origin (Madsen et al. 2010).
A mechanism enabling intercellular infection and formation of trans-cellular
infection threads within the root nodules was found to operate in the absence of
functional NFR1 and NFR5 receptors. Despite the independence from these
proteins known to be required for Nod-factor perception at early stages, the
formation of trans-cellular infection threads in nodules did depend on bacterial
production of intact Nod-factors, and nodC mutants of M. loti could not induce their
development (Madsen et al. 2010). It is yet unclear which molecules are involved in
the evident Nod-factor perception in the root cortex. Receptor kinases belonging to
the L. japonicus LysM receptor-like kinase (Lys) family, which includes NFR1 and
NFR5, are possible candidates (Lohmann et al. 2010). Of these, Lys genes that are
expressed predominantly in root and nodule tissues, namely Lys2, Lys3, Lys7,
Lys12, Lys15 and Lys20 (Lohmann et al. 2010), are the most likely to be involved.
A role of alternative epidermal or cortical Nod-factor receptors is further supported
Infection of Lotus japonicus Roots by Mesorhizobium loti
43
by observations in the tropical legume S. rostrata. Root hair infection was strictly
dependent on an intact Nod-factor structure, while root-hair independent crack
entry at lateral root bases was less stringently controlled in this species (D’Haeze
et al. 2000; Goormachtig et al. 2004).
The downstream components involved in signalling upon Nod-factor perception
by presumed cortical receptors are unknown. The complete absence of infection in
cyclops snf1 double mutants suggests that CCaMK activation followed by signalling through a CCaMK CYCLOPS complex is required for cortical infection thread
formation (Madsen et al. 2010). In contrast, infection studies in recently identified
mutants of the L. japonicus nucleoporin NENA suggest that crack entry and cortical
infection thread formation can occur independent of CCaMK activation through
Ca2+ spiking, or alternatively rely on rare activation events (Groth et al. 2010). In
line with the former interpretation, crack infection of the outer cortex was retained
in CCaMK knockdown-roots of Sesbania despite the loss of root hair infection and
nodulation (Capoen et al. 2009).
10
Alternative Infection Mechanisms: Single Cell Peg Entry
Characterisation of synthetic mutants combining gain of function (snf1) and loss of
function mutations lead to a remarkable and unexpected discovery of an infection
thread independent single cell entry mechanism in L. japonicus (Madsen et al.
2010). This intracellular infection of individual host cells by a process best
described as peg entry may constitute the ground state of bacterial invasion during
evolution of nodulation, and may allow for an analysis of the cardinal requirements
distinguishing endosymbiotic co-existence from pathogenesis. Surprisingly, this
entry mode was independent of both the NFR1 and NFR5 plant receptors and the
rhizobial Nod-factor signal molecule (Madsen et al. 2010). In a snf1 gain of
function genetic background, a bacterial nodC mutant unable to produce Nodfactors was capable of infecting nfr1 nfr5 single and double mutants although at a
20–100 fold reduced efficiency compared to normal root hair invasion. Infection
threads were not observed, but symbiosomes were present in the infected cells,
indicating that intercellular infection followed by endocytosis occurred in the
absence of Nod-factor and the NFR1 and NFR5 Nod-factor receptors (Madsen
et al. 2010). Such a capacity for direct intercellular infection could constitute the
ancient invasion path that evolved at the emergence of the legume family or
possibly at the emergence of the eurosid I clade containing all nodulating plants.
Continued division of the primary infected single cells would lead to the fully
infected Nod-factor independent nodulation observed in Aeschynomene type
nodules (Sprent 2007). Nod-factor independent stem nodulation of Aeschynomene
species (Giraud et al. 2007) appears to be a rare exception, however, and in many
legumes the subsequent evolutionary step may have been the Nod-factor and
cortical Nod-factor receptor dependent crack entry and subsequent cortical infection thread propagation of invasion as seen in S. rostrata (D’Haeze et al. 2000). The
44
K. Markmann et al.
Nod-factor dependent crack entry without infection threads and the fully infected
nodules in Arachis and Stylosanthes (Noti et al. 1985; Wilson et al. 1987; Boogerd
and van Rossum 1997) can be considered a variant of this infection mode. Fixation
thread symbiosis, where rhizobia are not released from the infection thread, as seen
in many legume trees and in Parasponia (Trinick 1979; Sprent 2007), is another
variant. Interestingly, the Gram-positive Frankia bacteria inducing actinorhizal
symbiosis in non-legumes are contained in fixation threads comparable to those
seen in legumes belonging to the oldest legume subfamily, the Caesalpinioideae
(Pawlowski and Bisseling 1996). Bacterial invasion via crack entry followed by the
formation of either fixation threads or infection threads with bacterial release into
symbiosomes in different members of the caesalpinoid genus Chamaecrista, support the notion of crack entry as a basal feature (Naisbitt et al. 1992; Sprent 2007).
The most highly evolved state suggested by the differential signalling and genetic
requirements of entry modes revealed in L. japonicus is root hair infection. This
infection mode required Nod-factor, epidermal NFR1 and NFR5 receptors, and is
predicted to also involve cortical Nod-factor receptors with less stringency than
NFR1 and NFR5. Overall this model predicts an evolutionary process leading from
pathogen-like bacterial invasion between cells to the more intimate containment
of – and coexistence with – symbiosomes inside plant cells.
Taken together, the advanced steps of bacterial release seen in the more recently
evolved legume hosts containing symbiosomes inside their cells have provided
these plants with a tighter and more selective control of bacterial proliferation and
nutrient exchange within the host. Similarly, the root hair infection mode seen in
many evolutionarily young legume groups allows for a tighter control of bacterial
passage through the epidermis than crack entry mechanisms. Observations of
infection modes in a wide selection of legumes belonging to different subfamilies
also indicate that alternative invasion modes have been maintained during evolution and that they are not mutually exclusive. Presence of different infection modes
in Sesbania (Goormachtig et al. 2004), Chamaecrista (Sprent 2007) and both
a Nod-factor dependent and a Nod-factor independent mechanism for nodulation
in Aeschynomene sensitiva (Giraud et al. 2007) is in accordance with this scenario.
Further comparison of the infection thread and direct infection pathway(s) seems to
provide a prime opportunity to assess the basal genetic components involved in the
release and endocytosis of bacteria.
References
Arrighi JF, Barre A, Ben Amor B, Bersoult A, Soriano LC, Mirabella R, de Carvalho-Niebel F,
Journet EP, Gherardi M, Huguet T, Geurts R, Denarie J, Rouge P, Gough C (2006) The
Medicago truncatula lysin motif-receptor-like kinase gene family includes NFP and new
nodule-expressed genes. Plant Physiol 142:265–279
Asamizu E, Shimoda Y, Kouchi H, Tabata S, Sato S (2008) A positive regulatory role for LjERF1
in the nodulation process is revealed by systematic analysis of nodule-associated transcription
factors of Lotus japonicus. Plant Physiol 147:2030–2040
Infection of Lotus japonicus Roots by Mesorhizobium loti
45
Banba M, Siddique AB, Kouchi H, Izui K, Hata S (2001) Lotus japonicus forms early senescent
root nodules with Rhizobium etli. Mol Plant Microbe Interact 14:173–180
Bek AS, Sauer J, Thygesen MB, Duus JO, Petersen BO, Thirup S, James E, Jensen KJ,
Stougaard J, Radutoiu S (2010) Improved characterization of nod factors and genetically
based variation in LysM Receptor domains identify amino acids expendable for nod factor
recognition in Lotus spp. Mol Plant Microbe Interact 23:58–66
Bohlool BB, Schmidt EL (1974) Lectins: a possible basis for specificity in the Rhizobium-legume
root nodule symbiosis. Science 185:269–271
Boogerd FC, van Rossum D (1997) Nodulation of groundnut by Bradyrhizobium: a simple
infection process by crack entry. FEMS Microbiol Rev 21:5–27
Borg S, Brandstrup B, Jensen TJ, Poulsen C (1997) Identification of new protein species among 33
different small GTP-binding proteins encoded by cDNAs from Lotus japonicus, and expression
of corresponding mRNAs in developing root nodules. Plant J 11:237–250
Bourne Y, Ayouba A, Rouge P, Cambillau C (1994) Interaction of a legume lectin with two
components of the bacterial cell wall. A crystallographic study. J Biol Chem 269:9429–9435
Brembu T, Winge P, Seem M, Bones AM (2004) NAPP and PIRP encode subunits of a putative wave regulatory protein complex involved in plant cell morphogenesis. Plant Cell 16:
2335–2349
Broughton WJ (2003) Roses by other names: taxonomy of the Rhizobiaceae. J Bacteriol 185:
2975–2979
Capoen W, Den Herder J, Sun J, Verplancke C, De Keyser A, De Rycke R, Goormachtig S,
Oldroyd G, Holsters M (2009) Calcium spiking patterns and the role of the calcium/calmodulin-dependent kinase CCaMK in lateral root base nodulation of Sesbania rostrata. Plant Cell
21:1526–1540
Cardenas L, Dominguez J, Quinto C, Lopez-Lara IM, Lugtenberg BJ, Spaink HP, Rademaker GJ,
Haverkamp J, Thomas-Oates JE (1995) Isolation, chemical structures and biological activity of
the lipo-chitin oligosaccharide nodulation signals from Rhizobium etli. Plant Mol Biol 29:
453–464
D’Antuono AL, Casabuono A, Couto A, Ugalde RA, Lepek VC (2005) Nodule development
induced by Mesorhizobium loti mutant strains affected in polysaccharide synthesis. Mol Plant
Microbe Interact 18:446–457
D’Antuono AL, Ott T, Krusell L, Voroshilova V, Ugalde RA, Udvardi M, Lepek VC (2008)
Defects in rhizobial cyclic glucan and lipopolysaccharide synthesis alter legume gene expression during nodule development. Mol Plant Microbe Interact 21:50–60
De Hoff PL, Brill LM, Hirsch AM (2009) Plant lectins: the ties that bind in root symbiosis and
plant defense. Mol Genet Genomics 282:1–15
Deeks MJ, Kaloriti D, Davies B, Malho R, Hussey PJ (2004) Arabidopsis NAP1 is essential for
Arp2/3-dependent trichome morphogenesis. Curr Biol 14:1410–1414
D’Haeze W, Mergaert P, Prome JC, Holsters M (2000) Nod factor requirements for efficient stem
and root nodulation of the tropical legume Sesbania rostrata. J Biol Chem 275:15676–15684
Fabra A, Castro S, Taurian T, Angelini J, Ibanez F, Dardanelli M, Tonelli M, Bianucci E, Valetti L
(2010) Interaction among Arachis hypogaea L. (peanut) and beneficial soil microorganisms:
how much is it known? Crit Rev Microbiol 36:179–194
Gage DJ (2004) Infection and invasion of roots by symbiotic, nitrogen-fixing rhizobia during
nodulation of temperate legumes. Microbiol Mol Biol Rev 68:280–300
Giraud E, Moulin L, Vallenet D, Barbe V, Cytryn E, Avarre JC, Jaubert M, Simon D, Cartieaux F,
Prin Y, Bena G, Hannibal L, Fardoux J, Kojadinovic M, Vuillet L, Lajus A, Cruveiller S,
Rouy Z, Mangenot S, Segurens B, Dossat C, Franck WL, Chang WS, Saunders E, Bruce D,
Richardson P, Normand P, Dreyfus B, Pignol D, Stacey G, Emerich D, Vermeglio A, Medigue
C, Sadowsky M (2007) Legumes symbioses: absence of Nod genes in photosynthetic
bradyrhizobia. Science 316:1307–1312
Goormachtig S, Capoen W, James E, Holsters M (2004) Switch from intracellular to intercellular
invasion during water stress-tolerant legume nodulation. Proc Natl Acad Sci USA 101:
6303–6308
46
K. Markmann et al.
Groth M, Takeda N, Perry J, Uchida H, Draxl S, Brachmann A, Sato S, Tabata S, Kawaguchi M,
Wang TL, Parniske M (2010) NENA, a Lotus japonicus homolog of Sec13, is required for
rhizodermal infection by arbuscular mycorrhiza fungi and rhizobia but dispensable for cortical
endosymbiotic development. Plant Cell 22:2509–2526
Haney CH, Long SR (2010) Plant flotillins are required for infection by nitrogen-fixing bacteria.
Proc Natl Acad Sci USA 107:478–483
Hayashi T, Banba M, Shimoda Y, Kouchi H, Hayashi M, Imaizumi-Anraku H (2010) A dominant
function of CCaMK in intracellular accommodation of bacterial and fungal endosymbionts.
Plant J 63:141–154
Heckmann AB, Lombardo F, Miwa H, Perry JA, Bunnewell S, Parniske M, Wang TL, Downie JA
(2006) Lotus japonicus nodulation requires two GRAS domain regulators, one of which is
functionally conserved in a non-legume. Plant Physiol 142:1739–1750
Held M, Hossain MS, Yokota K, Bonfante P, Stougaard J, Szczyglowski K (2010) Common and
not so common symbiotic entry. Trends Plant Sci 15:540–545
Hirsch S, Kim J, Munoz A, Heckmann AB, Downie JA, Oldroyd GE (2009) GRAS proteins form a
DNA binding complex to induce gene expression during nodulation signaling in Medicago
truncatula. Plant Cell 21:545–557
Hoegslund N, Radutoiu S, Krusell L, Voroshilova V, Hannah MA, Goffard N, Sanchez DH,
Lippold F, Ott T, Sato S, Tabata S, Liboriussen P, Lohmann GV, Schauser L, Weiller GF,
Udvardi MK, Stougaard J (2009) Dissection of symbiosis and organ development by integrated
transcriptome analysis of Lotus japonicus mutant and wild-type plants. PLoS One 4:e6556
Hotter GS, Scott DB (1991) Exopolysaccharide mutants of Rhizobium loti are fully effective on a
determinate nodulating host but are ineffective on an indeterminate nodulating host. J Bacteriol
173:851–859
Hubber A, Vergunst AC, Sullivan JT, Hooykaas PJ, Ronson CW (2004) Symbiotic phenotypes and
translocated effector proteins of the Mesorhizobium loti strain R7A VirB/D4 type IV secretion
system. Mol Microbiol 54:561–574
Hubber AM, Sullivan JT, Ronson CW (2007) Symbiosis-induced cascade regulation of the
Mesorhizobium loti R7A VirB/D4 type IV secretion system. Mol Plant Microbe Interact 20:
255–261
Hussain AKMA, Jiang Q, Broughton WJ, Gresshoff PM (1999) Lotus japonicus nodulates and
fixes nitrogen with the broad host range Rhizobium sp. NGR234. Plant Cell Physiol 40:
894–899
Imaizumi-Anraku H, Kawaguchi M, Koiwa H, Akao S, Syono K (1997) Two ineffectivenodulating mutants of Lotus japonicus – different phenotypes caused by the blockage of
endocytotic bacterial release and nodule maturation. Plant Cell Physiol 38:871–881
Imaizumi-Anraku H, Kouchi H, Syono K, Akao S, Kawaguchi M (2000) Analysis of ENOD40
expression in alb1, a symbiotic mutant of Lotus japonicus that forms empty nodules with
incompletely developed nodule vascular bundles. Mol Gen Genet 264:402–410
Imaizumi-Anraku H, Takeda N, Charpentier M, Perry J, Miwa H, Umehara Y, Kouchi H,
Murakami Y, Mulder L, Vickers K, Pike J, Downie JA, Wang T, Sato S, Asamizu E, Tabata
S, Yoshikawa M, Murooka Y, Wu GJ, Kawaguchi M, Kawasaki S, Parniske M, Hayashi M
(2005) Plastid proteins crucial for symbiotic fungal and bacterial entry into plant roots. Nature
433:527–531
Kanamori N, Madsen LH, Radutoiu S, Frantescu M, Quistgaard EM, Miwa H, Downie JA, James
EK, Felle HH, Haaning LL, Jensen TH, Sato S, Nakamura Y, Tabata S, Sandal N, Stougaard J
(2006) A nucleoporin is required for induction of Ca2+ spiking in legume nodule development
and essential for rhizobial and fungal symbiosis. Proc Natl Acad Sci USA 103:359–364
Karas B, Murray J, Gorzelak M, Smith A, Sato S, Tabata S, Szczyglowski K (2005) Invasion of
Lotus japonicus root hairless 1 by Mesorhizobium loti involves the nodulation factor-dependent induction of root hairs. Plant Physiol 137:1331–1344
Lefebvre B, Timmers T, Mbengue M, Moreau S, Herve C, Toth K, Bittencourt-Silvestre J,
Klaus D, Deslandes L, Godiard L, Murray JD, Udvardi MK, Raffaele S, Mongrand S,
Infection of Lotus japonicus Roots by Mesorhizobium loti
47
Cullimore J, Gamas P, Niebel A, Ott T (2010) A remorin protein interacts with symbiotic
receptors and regulates bacterial infection. Proc Natl Acad Sci USA 107:2343–2348
Lerouge P, Roche P, Faucher C, Maillet F, Truchet G, Prome JC, Denarie J (1990) Symbiotic hostspecificity of Rhizobium meliloti is determined by a sulphated and acylated glucosamine
oligosaccharide signal. Nature 344:781–784
Li Y, Sorefan K, Hemmann G, Bevan MW (2004) Arabidopsis NAP and PIR regulate actin-based
cell morphogenesis and multiple developmental processes. Plant Physiol 136:3616–3627
Limpens E, Ivanov S, van Esse W, Voets G, Fedorova E, Bisseling T (2009) Medicago N2-fixing
symbiosomes acquire the endocytic identity marker Rab7 but delay the acquisition of vacuolar
identity. Plant Cell 21:2811–2828
Lohar DP, Schaff JE, Laskey JG, Kieber JJ, Bilyeu KD, Bird DM (2004) Cytokinins play opposite
roles in lateral root formation, and nematode and Rhizobial symbioses. Plant J 38:203–214
Lohmann GV, Shimoda Y, Nielsen MW, Jorgensen FG, Grossmann C, Sandal N, Sorensen K,
Thirup S, Madsen LH, Tabata S, Sato S, Stougaard J, Radutoiu S (2010) Evolution and
regulation of the Lotus japonicus LysM receptor gene family. Mol Plant Microbe Interact
23:510–521
Lombardo F, Heckmann AB, Miwa H, Perry JA, Yano K, Hayashi M, Parniske M, Wang TL,
Downie JA (2006) Identification of symbiotically defective mutants of Lotus japonicus
affected in infection thread growth. Mol Plant Microbe Interact 19:1444–1450
Lopez-Lara IM, van den Berg JD, Thomas-Oates JE, Glushka J, Lugtenberg BJ, Spaink HP (1995)
Structural identification of the lipo-chitin oligosaccharide nodulation signals of Rhizobium loti.
Mol Microbiol 15:627–638
Madsen EB, Madsen LH, Radutoiu S, Olbryt M, Rakwalska M, Szczyglowski K, Sato S,
Kaneko T, Tabata S, Sandal N, Stougaard J (2003) A receptor kinase gene of the LysM type
is involved in legume perception of rhizobial signals. Nature 425:637–640
Madsen LH, Tirichine L, Jurkiewicz A, Sullivan JT, Heckmann AB, Bek AS, Ronson CW,
James EK, Stougaard J (2010) The molecular network governing nodule organogenesis and
infection in the model legume Lotus japonicus. Nat Commun 1:1–12
Madsen EB, Antolin-Llovera M, Grossmann C, Ye J, Vieweg S, Broghammer A, Krusell L,
Radutoiu S, Jensen ON, Stougaard J, Parniske M (2011) Autophosphorylation is essential for
the in vivo function of the Lotus japonicus Nod factor receptor 1 and receptor-mediated
signalling in cooperation with Nod factor receptor 5. Plant J 65:404–417
Maillet F, Poinsot V, Andre O, Puech-Pages V, Haouy A, Gueunier M, Cromer L, Giraudet D,
Formey D, Niebel A, Martinez EA, Driguez H, Becard G, Denarie J (2011) Fungal lipochitooligosaccharide symbiotic signals in arbuscular mycorrhiza. Nature 469:58–63
Messinese E, Mun JH, Yeun LH, Jayaraman D, Rouge P, Barre A, Lougnon G, Schornack S,
Bono JJ, Cook DR, Ane JM (2007) A novel nuclear protein interacts with the symbiotic DMI3
calcium- and calmodulin-dependent protein kinase of Medicago truncatula. Mol Plant
Microbe Interact 20:912–921
Middleton PH, Jakab J, Penmetsa RV, Starker CG, Doll J, Kalo P, Prabhu R, Marsh JF, Mitra RM,
Kereszt A, Dudas B, VandenBosch K, Long SR, Cook DR, Kiss GB, Oldroyd GE (2007) An
ERF transcription factor in Medicago truncatula that is essential for Nod factor signal
transduction. Plant Cell 19:1221–1234
Miwa H, Sun J, Oldroyd GE, Downie JA (2006) Analysis of Nod-factor-induced calcium signaling
in root hairs of symbiotically defective mutants of Lotus japonicus. Mol Plant Microbe Interact
19:914–923
Miya A, Albert P, Shinya T, Desaki Y, Ichimura K, Shirasu K, Narusaka Y, Kawakami N, Kaku H,
Shibuya N (2007) CERK1, a LysM receptor kinase, is essential for chitin elicitor signaling in
Arabidopsis. Proc Natl Acad Sci USA 104:19613–19618
Moulin L, Munive A, Dreyfus B, Boivin-Masson C (2001) Nodulation of legumes by members of
the beta-subclass of Proteobacteria. Nature 411:948–950
Mulligan JT, Long SR (1985) Induction of Rhizobium meliloti nodC expression by plant exudate
requires nodD. Proc Natl Acad Sci USA 82:6609–6613
48
K. Markmann et al.
Murakami EI, Nagata M, Shimoda Y, Kucho KI, Higashi S, Abe M, Hashimoto M, Uchiumi T
(2011) Nitric oxide production induced in roots of Lotus japonicus by lipopolysaccharide from
Mesorhizobium loti. Plant Cell Physiol 52:610–617
Murray JD, Karas BJ, Sato S, Tabata S, Amyot L, Szczyglowski K (2007) A cytokinin perception
mutant colonized by Rhizobium in the absence of nodule organogenesis. Science 315:101–104
Naisbitt T, James EK, Sprent JI (1992) The evolutionary significance of the legume genus
Chamaecrista, as determined by nodule structure. New Phytol 122:487–492
Nakagawa T, Kaku H, Shimoda Y, Sugiyama A, Shimamura M, Takanashi K, Yazaki K, Aoki T,
Shibuya N, Kouchi H (2011) From defense to symbiosis: limited alterations in the kinase
domain of LysM receptor-like kinases are crucial for evolution of legume-Rhizobium symbiosis. Plant J 65:169–180
Noti JD, Dudas B, Szalay AA (1985) Isolation and characterization of nodulation genes from
Bradyrhizobium sp. (Vigna) strain IRc 78. Proc Natl Acad Sci USA 82:7379–7383
Okazaki S, Okabe S, Higashi M, Shimoda Y, Sato S, Tabata S, Hashiguchi M, Akashi R,
Gottfert M, Saeki K (2010) Identification and functional analysis of type III effector proteins
in Mesorhizobium loti. Mol Plant Microbe Interact 23:223–234
Oldroyd GED, Downie JA (2004) Calcium, kinases and nodulation signalling in legumes. Nat Rev
Mol Cell Biol 5:566–576
Pacios-Bras C (2003). The symbiosis between Lotus japonicus and rhizobia: Function of nod
factor structural variation. PhD Thesis, University of Leiden, Leiden, The Netherlands
Pawlowski K, Bisseling T (1996) Rhizobial and actinorhizal symbioses: what are the shared
features? Plant Cell 8:1899–1913
Peters NK, Frost JW, Long SR (1986) A plant flavone, luteolin, induces expression of Rhizobium
meliloti nodulation genes. Science 233:977–980
Radutoiu S, Madsen LH, Madsen EB, Felle HH, Umehara Y, Groenlund M, Sato S, Nakamura Y,
Tabata S, Sandal N, Stougaard J (2003) Plant recognition of symbiotic bacteria requires two
LysM receptor-like kinases. Nature 425:585–592
Radutoiu S, Madsen LH, Madsen EB, Jurkiewicz A, Fukai E, Quistgaard EM, Albrektsen AS,
James EK, Thirup S, Stougaard J (2007) LysM domains mediate lipochitin-oligosaccharide
recognition and Nfr genes extend the symbiotic host range. EMBO J 26:3923–3935
Rodpothong P, Sullivan JT, Songsrirote K, Sumpton D, Cheung KW, Thomas-Oates J, Radutoiu S,
Stougaard J, Ronson CW (2009) Nodulation gene mutants of Mesorhizobium loti R7A-nodZ
and nolL mutants have host-specific phenotypes on Lotus spp. Mol Plant Microbe Interact 22:
1546–1554
Saito K, Yoshikawa M, Yano K, Miwa H, Uchida H, Asamizu E, Sato S, Tabata S, ImaizumiAnraku H, Umehara Y, Kouchi H, Murooka Y, Szczyglowski K, Downie JA, Parniske M,
Hayashi M, Kawaguchi M (2007) NUCLEOPORIN85 is required for calcium spiking, fungal
and bacterial symbioses, and seed production in Lotus japonicus. Plant Cell 19:610–624
Sanchez C, Iannino F, Deakin WJ, Ugalde RA, Lepek VC (2009) Characterization of the
Mesorhizobium loti MAFF303099 type-three protein secretion system. Mol Plant Microbe
Interact 22:519–528
Schauser L, Handberg K, Sandal N, Stiller J, Thykjaer T, Pajuelo E, Nielsen A, Stougaard J (1998)
Symbiotic mutants deficient in nodule establishment identified after T-DNA transformation of
Lotus japonicus. Mol Gen Genet 259:414–423
Schauser L, Roussis A, Stiller J, Stougaard J (1999) A plant regulator controlling development of
symbiotic root nodules. Nature 402:191–195
Schumpp O, Crevecoeur M, Broughton WJ, Deakin WJ (2009) Delayed maturation of nodules
reduces symbiotic effectiveness of the Lotus japonicus-Rhizobium sp. NGR234 interaction.
J Exp Bot 60:581–590
Spaink HP, Okker RJ, Wijffelman CA, Tak T, Goosen-de Roo L, Pees E, van Brussel AA,
Lugtenberg BJ (1989) Symbiotic properties of rhizobia containing a flavonoid-independent
hybrid nodD product. J Bacteriol 171:4045–4053
Infection of Lotus japonicus Roots by Mesorhizobium loti
49
Spaink HP, Sheeley DM, van Brussel AA, Glushka J, York WS, Tak T, Geiger O, Kennedy EP,
Reinhold VN, Lugtenberg BJ (1991) A novel highly unsaturated fatty acid moiety of lipooligosaccharide signals determines host specificity of Rhizobium. Nature 354:125–130
Sprent JI (2007) Evolving ideas of legume evolution and diversity: a taxonomic perspective on the
occurrence of nodulation. New Phytol 174:11–25
Stracke S, Kistner C, Yoshida S, Mulder L, Sato S, Kaneko T, Tabata S, Sandal N, Stougaard J,
Szczyglowski K, Parniske M (2002) A plant receptor-like kinase required for both bacterial
and fungal symbiosis. Nature 417:959–962
Szczyglowski K, Shaw RS, Wopereis J, Copeland S, Hamburger D, Kasiborski B, Dazzo FB, de
Bruijn FJ (1998) Nodule organogenesis and symbiotic mutants of the model legume Lotus
japonicus. Mol Plant Microb Interact 11:684–697
Tansengco ML, Hayashi M, Kawaguchi M, Imaizumi-Anraku H, Murooka Y (2003) crinkle, a
novel symbiotic mutant that affects the infection thread growth and alters the root hair,
trichome, and seed development in Lotus japonicus. Plant Physiol 131:1054–1063
Tirichine L, Imaizumi-Anraku H, Yoshida S, Murakami Y, Madsen LH, Miwa H, Nakagawa T,
Sandal N, Albrektsen AS, Kawaguchi M, Downie A, Sato S, Tabata S, Kouchi H, Parniske M,
Kawasaki S, Stougaard J (2006) Deregulation of a Ca2+/calmodulin-dependent kinase leads to
spontaneous nodule development. Nature 441:1153–1156
Tirichine L, Sandal N, Madsen LH, Radutoiu S, Albrektsen AS, Sato S, Asamizu E, Tabata S,
Stougaard J (2007) A gain-of-function mutation in a cytokinin receptor triggers spontaneous
root nodule organogenesis. Science 315:104–107
Trinick MJ (1979) Structure of nitrogen-fixing nodules formed by Rhizobium on roots of
Parasponia andersonii Planch. Can J Microbiol 25:565–578
Truchet G, Roche P, Lerouge P, Vasse J, Camut S, de Billy F, Prome J-C, Denarie J (1991)
Sulphated lipo-oligosaccharide signals of Rhizobium meliloti elicit root nodule organogenesis
in alfalfa. Nature 351:670–673
Van de Velde W, Zehirov G, Szatmari A, Debreczeny M, Ishihara H, Kevei Z, Farkas A, Mikulass
K, Nagy A, Tiricz H, Satiat-Jeunemaitre B, Alunni B, Bourge M, Kucho K, Abe M, Kereszt A,
Maroti G, Uchiumi T, Kondorosi E, Mergaert P (2010) Plant peptides govern terminal
differentiation of bacteria in symbiosis. Science 327:1122–1126
van Rhijn P, Goldberg RB, Hirsch AM (1998) Lotus corniculatus nodulation specificity is changed
by the presence of a soybean lectin gene. Plant Cell 10:1233–1250
van Rhijn P, Fujishige NA, Lim PO, Hirsch AM (2001) Sugar-binding activity of pea lectin
enhances heterologous infection of transgenic alfalfa plants by Rhizobium leguminosarum
biovar viciae. Plant Physiol 126:133–144
van Spronsen PC, Groenlund M, Bras CP, Spaink HP, Kijne JW (2001) Cell biological changes of
outer cortical root cells in early determinate nodulation. Mol Plant Microbe Interact 14:
839–847
Vassileva VN, Kouchi H, Ridge RW (2005) Microtubule dynamics in living root hairs: transient
slowing by lipochitin oligosaccharide nodulation signals. Plant Cell 17:1777–1787
Wan J, Zhang XC, Neece D, Ramonell KM, Clough S, Kim SY, Stacey MG, Stacey G (2008) A
LysM receptor-like kinase plays a critical role in chitin signaling and fungal resistance in
Arabidopsis. Plant Cell 20:471–481
Weerasinghe RR, Collings DA, Johannes E, Allen NS (2003) The distributional changes and role
of microtubules in Nod factor-challenged Medicago sativa root hairs. Planta 218:276–287
Weerasinghe RR, Bird DM, Allen NS (2005) Root-knot nematodes and bacterial Nod factors
elicit common signal transduction events in Lotus japonicus. Proc Natl Acad Sci USA 102:
3147–3152
Wilson KJ, Anjaiah V, Nambiar PT, Ausubel FM (1987) Isolation and characterization of
symbiotic mutants of Bradyrhizobium sp. (Arachis) strain NC92: mutants with host-specific
defects in nodulation and nitrogen fixation. J Bacteriol 169:2177–2186
Yang S, Tang F, Gao M, Krishnan HB, Zhu H (2010) R gene-controlled host specificity in the
legume-rhizobia symbiosis. Proc Natl Acad Sci USA 107:18735–18740
50
K. Markmann et al.
Yano K, Tansengco ML, Hio T, Higashi K, Murooka Y, Imaizumi-Anraku H, Kawaguchi M,
Hayashi M (2006) New nodulation mutants responsible for infection thread development in
Lotus japonicus. Mol Plant Microbe Interact 19:801–810
Yano K, Yoshida S, Muller J, Singh S, Banba M, Vickers K, Markmann K, White C, Schuller B,
Sato S, Asamizu E, Tabata S, Murooka Y, Perry J, Wang TL, Kawaguchi M, Imaizumi-Anraku
H, Hayashi M, Parniske M (2008) CYCLOPS, a mediator of symbiotic intracellular accommodation. Proc Natl Acad Sci USA 105:20540–20545
Yano K, Shibata S, Chen WL, Sato S, Kaneko T, Jurkiewicz A, Sandal N, Banba M, ImaizumiAnraku H, Kojima T, Ohtomo R, Szczyglowski K, Stougaard J, Tabata S, Hayashi M,
Kouchi H, Umehara Y (2009) CERBERUS, a novel U-box protein containing WD-40 repeats,
is required for formation of the infection thread and nodule development in the legumeRhizobium symbiosis. Plant J 60:168–180
Yokota K, Fukai E, Madsen LH, Jurkiewicz A, Rueda P, Radutoiu S, Held M, Hossain MS,
Szczyglowski K, Morieri G, Oldroyd GE, Downie JA, Nielsen MW, Rusek AM, Sato S,
Tabata S, James EK, Oyaizu H, Sandal N, Stougaard J (2009) Rearrangement of actin
cytoskeleton mediates invasion of Lotus japonicus roots by Mesorhizobium loti. Plant Cell
21:267–284
Zhukov V, Radutoiu S, Madsen LH, Rychagova T, Ovchinnikova E, Borisov A, Tikhonovich I,
Stougaard J (2008) The pea Sym37 receptor kinase gene controls infection-thread initiation and
nodule development. Mol Plant Microbe Interact 21:1600–1608
Signalling and the Re-structuring of Plant
Cell Architecture in AM Symbiosis
Andrea Genre
Abstract Arbuscular mycorrhizas are widespread and ancient plant symbioses that
were already established by the first land plants when they abandoned the water
environment. Arbuscular mycorrhizal fungi scavenge mineral nutrients and water
from the soil improving the overall plant fitness, receiving in exchange
carbohydrates that are indispensable to complete their life cycle. In spite of their
importance in natural and agricultural ecosystems, many biological aspects of these
interactions are still partially obscure, especially concerning the early stages of
symbiosis establishment which involve a signal exchange between the partners.
Nonetheless, recent advancements have started to shed light on plant–fungus
signalling mechanisms and their relation with the cell responses that culminate in
fungal accommodation in the root cells. This chapter is focused on such advances
and the new views that they have suggested.
1 Introduction
Arbuscular mycorrhizas (AM) are considered to be the most ancient plant symbiosis. Fossil data report the presence of arbuscules inside the tissues of some of the
first plants that abandoned the water environment to start dry land colonisation
(Remy et al. 1994). The ability to scavenge mineral nutrients and water from humid
soils has likely been the key to the success of AM fungi, and hence of the plants that
were hosting them. Present-day soils retain their nutrients as strongly as those
prehistoric shores (Leeper 1952). It is therefore not surprising that AM interactions
are still so popular in the plant kingdom. Eighty to ninety percent of all plant species
interact in natural and agronomical environments with AM fungi (Smith and Read
2008).
A. Genre (*)
Dipartimento di Biologia Vegetale, Università di Torino, Viale Mattioli 25, 10125 Torino, Italy
e-mail: [email protected]
S. Perotto and F. Baluska (eds.), Signaling and Communication in Plant Symbiosis,
Signaling and Communication in Plants 11, DOI 10.1007/978-3-642-20966-6_3,
# Springer-Verlag Berlin Heidelberg 2012
51
52
A. Genre
In spite of its widespread distribution and basic importance in plant nutrition, our
knowledge of several biological aspects of the AM symbiosis is still patchy,
especially when compared to other plant interactions, such as those with pathogenic
fungi or oomycetes. Beside the economical aspects (defeating a pathogen has been
so far much more profitable than exchanging chemical fertilisers for symbiotic
fungi), several biological reasons account for this knowledge gap. First of all, AM
fungi, which all belong to the small phylum of Glomeromycota, are rather unique
organisms, probably also as a consequence of their long co-evolution with plants.
AM fungi cannot reproduce in the absence of their hosts, they are strictly asexual
and their huge spores (up to 500 mm) may contain a mixed population of thousands
of nuclei, bearing different sets of genes (Hijri and Sanders 2005; Sanders and Croll
2010). The very concepts of individual, species and cell are difficult to apply to
these fungi (Pawlowska and Taylor 2004), which are aseptate and even anastomose
with each other mixing their respective nuclear populations through strong cytoplasmic streams (Croll et al. 2009). Summing up so many unusual features, AM
fungi clearly do not fit the definition of a model organism for laboratory
investigations. For example, classic genetic studies are totally unsuitable for these
fungi (Vandenkoornhuyse et al. 2001). On top of that, rather than developing on a
leaf – in full view of an attentive observer – AM interactions take place underneath
the soil surface, strongly hampering our chances to understand most of the early
steps of their development, including signal exchanges between the plant and
fungus. These have only recently been approached thanks to important technical
advancements, such as the ability to obtain and follow the symbiosis development
inside a Petri dish (Bécard and Fortin 1988; Cano et al. 2008), and have rapidly
attained the mainstream of AM research.
After a quick overview of the root colonization process by AM fungi, this
chapter will focus on the recent advancements in the study of plant–fungus signal
exchanges and their relation with the plant responses that culminate in fungal
accommodation inside the root cells. Compared to other plant–microbe
interactions, the resulting picture is still missing many major elements, but new
chances can now be taken to peek into the biological processes that control the
establishment of AM symbiosis.
2 The Process of Root Colonization
The development of root colonization by Glomeromycota is largely conserved
throughout host plant taxa. Under favourable environmental conditions, the large
asexual spores of AM fungi produce germination hyphae that can only grow for a
few days in the absence of a host plant (asymbiotic phase). This short mycelium is
only fed by the spore, as it is unable to uptake carbon from the soil organic matter
(Leight et al. 2009), and its life span is limited to a few days, after which the hyphae
stop growing and retract their cytoplasm into the spore, in view of a new germination event. By contrast, as soon as AM hyphae reach the vicinity of a host root
Signalling and the Re-structuring of Plant Cell Architecture in AM Symbiosis
53
(presymbiotic phase), they visibly change their developmental program and branch
repeatedly (Buee et al. 2000). This phase, when the plant and fungus perceive each
other, is followed by direct plant–fungus contact, which associates with the setup of
novel developmental and cellular modifications in both partners. The hyphae of
glomeromycetes normally explore a stretch of the contacted root before changing
their growth pattern to form highly branched, swollen and flattened hyphal
protrusions that strongly adhere to the wall of epidermal cells (Smith and Read
2008). Such a structure is known as a hyphopodium (Bastmeyer et al. 2002) and its
adhesion to the atrichoblasts of the root epidermis marks the initiation of the
symbiotic phase of the interaction. Root epidermal cells in turn respond to
hyphopodium contact by repositioning their nucleus and remodelling their cytoplasm, in preparation for fungal penetration (Genre et al. 2005). In fact, the entry of
AM fungi in the plant cell lumen implies the development of a novel cellular
compartment, the symbiotic interface. This structure, of plant origin, is composed
of a thin layer of cell wall materials in direct contact with the fungus, and an
invagination of the host plasma membrane – named the perifungal membrane
(Bonfante 2001). The interface compartment envelopes every intracellular hypha,
from the simplest, straight hypha crossing an epidermal cell to the highly branched
arbuscule filling up most of a cortical cell volume. In most cases fungal colonisation
proceeds intracellularly across the epidermal and outer cortical cells. By contrast,
once the hyphal tips have reached the inner cortex they can also develop along the
intercellular spaces of the cortical parenchyma (Dickson 2004). This intercellular
mode of growth is particularly common in legumes, and in general in those plants
(e.g. Arum sp.) whose root anatomy presents such extensive apoplastic channels. In
these cases, most of the arbuscules develop from terminal hyphal branches, whereas
files of intercalary arbuscules originate when inner cortex colonisation mostly
proceeds from a cell to the next with a so-called intracellular mode of growth
(e.g. carrot or Paris sp.).
Arbuscules – the structures that give AM interactions their name – originate
from the repeated branching of an intracellular hypha. Arbusculated cells are the
site where symbiosis achieves its highest degree of complexity from a morphological and functional point of view. The symbiotic interface, lined by the
periarbuscular membrane, envelopes each fine branch of the arbuscule and
mediates nutrient – and likely signal – exchanges. Originally suggested by the
very high surface-to-volume ratio achieved by the arbuscule recursive branching
(Dickson and Kolesik 1999), the concentration of nutrient exchange in the
arbuscules has recently found more direct evidence in the identification of plant
phosphate and ammonium transporters that are mainly expressed in arbusculated
cells (Javot et al. 2007; Balestrini et al. 2007; Guether et al. 2009) and localized on
the periarbuscular membrane (Pumplin and Harrison 2009; Kobae and Hata 2010).
The establishment of a functional AM symbiosis therefore involves a progressive increase in the closeness of the interaction, from the exchange of long-range
chemical signals in the rhizosphere to intimate intracellular association, when the
plant and fungus share a single cell volume.
54
A. Genre
3 The Presymbiotic Dialogue
Increasing evidence has accumulated over the years concerning the exchange of
chemical signals between the plant and fungus in advance of their first physical
contact. A summary of the resulting picture is presented in Fig. 1. Keeping both
partners informed on their proximity is an obvious advantage in preparation for
direct interaction. In the case of AM, this is achieved through an apparently rough,
but successful strategy: the perception of plant diffusible signals induces hyphal
ramification within hours (Akiyama et al. 2005), while fungal proximity stimulates
lateral root development in days (Olah et al. 2005). As a consequence, the chances
for the hyphae and roots to eventually meet increase exponentially with reciprocal
vicinity. Such presymbiotic developmental responses associate with metabolic,
transcriptional and cellular responses, as part of an “anticipation program” that
prepares both symbionts to a successful association (Paszkowski 2006a).
Fig. 1 Presymbiotic signalling. The scheme summarises the main responses observed in both the
plant and fungus during the presymbiotic phase of an AM interaction. Strigolactones released by
the host plant root activate fungal metabolism, stimulate hyphal branching and induce gene
regulation. At the same time, the perception of a so-far unidentified Myc factor by the plant is
mediated by the SYM transduction pathway and leads to gene regulation, starch accumulation in
the root and the development of new lateral roots
Signalling and the Re-structuring of Plant Cell Architecture in AM Symbiosis
3.1
55
Strigolactones: Eclectic Plant Messengers
Direct evidence for a chemotropic development has only been found for a few
glomeromycetes (Sbrana and Giovannetti 2005), while Akiyama et al. (2005) have
identified a strigolactone secreted by Lotus japonicus roots as the molecule responsible for stimulating such host-induced hyphal branching in AM fungi. Furthermore, Sorghum strigolactones have also been shown to boost AM hyphal growth
and metabolism, and increase mitochondrial number and motility (Besserer and
Puech-Pagès 2006; Besserer et al. 2008). Strigolactone instability in water and their
continuous release from the root combine into a steep concentration gradient, which
provides precise topological information about root localization and directly
impacts the pattern of fungal development.
Interestingly, the biological activity of strigolactones goes beyond AM
interactions, as confirmed by the fact that their production is not limited to AM
host plants (Delaux et al. 2009): such carotenoid-derived molecules have in fact
several and very diverse functions. Their principal role is likely to be hormonal.
Being produced both in the root and shoot, strigolactones control shoot branching
by inhibiting lateral bud development (Gómez-Roldán et al. 2008). Their role as
rhizospheric messengers seems therefore to have originated secondarily, due to
their continuous leakage from the roots. Not surprisingly, the strigolactone gradient
is also exploited by plant parasites. Seed germination in parasitic plants such as Striga
(Cook et al. 1966) is boosted by the presence of root-exudated strigolactones – whose
name in fact derives from this very plant genus. Tomato plants bearing a mutation
in a gene involved in strigolactone biosynthesis (SlORT1) are both resistant to the
parasitic weed Orobanche and reluctant to AM colonization (Koltai et al. 2010).
Nevertheless, Akiyama et al. (2010) have recently highlighted the fine structural
requirements that specifically correlate with the activity of strigolactones in
stimulating AM fungal branching versus parasitic plant seed germination. It therefore appears likely that, during the co-evolution of different organisms with plants,
differential signalling mechanisms have tuned to specific forms of strigolactones.
In spite of the long studies carried out on parasitic plants, and the more recent
investigations on AM fungi and epigeous plant tissues, one missing tile in this
complex communication system is the identification of the proteins that mediate
strigolactone perception. Two genes encoding for an F-Box protein and an a/b-fold
hydrolase have recently been identified as potential strigolactone receptors in the
shoot of rice (Ishikawa et al. 2005; Arite et al. 2009), and a 60 kDa strigolactonebinding protein has been isolated from Striga hermontica (Zwanenburg et al. 2009).
Such studies may open the way to the identification of proteins with an analogous
function in the fungus.
In conclusion, strigolactones are considered as the main actors in the plant-tofungus communications that precede their physical interaction. We cannot anyway
exclude that other factors are also involved. If we consider the relatively similar
case of nitrogen-fixing root nodules, beside the acknowledged flavonoid-based
signalling mechanism required for symbiosis establishment, a role has recently
56
A. Genre
been suggested for strigolactones also (Soto et al. 2010), thus introducing a possible
crosstalk between the early events that control legume root symbioses.
3.2
The Elusive AM Fungal Signals
The perception of fungal diffusible signals by the host plants was demonstrated by
several publications over the last years. Such signals are often referred to as “Myc
factor” in analogy to the Nod factor produced by nitrogen fixing rhizobia (Kosuta
et al. 2003), Maillet et al. (2011). Myc factor perception induces the first molecular
and cellular responses in the host plants several hours before the development of
hyphopodia on the root surface. The Myc factor diffuses in water solutions and can
cross permeable membranes that prevent physical contacts between the two
partners. In these conditions, fungal diffusible factors were shown to activate the
expression of the early nodulin MtEnod11 and other symbiosis-related genes that
are also expressed during root colonization (Kosuta et al. 2003; Weidmann et al.
2004). More recently, a membrane steroid binding protein (MtMSBP1) and other
plant genes have also been convincingly shown to be upregulated in the root tissues
approached by the branching AM hyphae (Kuhn et al. 2010).
Beside the stimulation of lateral root development (Olah et al. 2005), another
long-term host plant response to diffusible fungal signals was detected while
investigating root starch content. A decrease in root starch level is normally
associated with arbuscule development. By contrast, when L. japonicus plants
were exposed for 1 week to AM fungal exudates or grown in the presence of a
membrane to prevent fungal contact and colonization, starch accumulated in the
root cortex and was stored there for at least 1 month (Gutjahr et al. 2009). Such a
change in carbohydrate mobilisation can easily be interpreted as another element of
the anticipation program, preparing the carbohydrate supplies that should normally
be bound for the fungus.
Such studies demonstrate beyond question that host plants respond to diffusible
AM fungal signals. Nonetheless, many aspects of the responsible signalling
mechanisms remain unknown. First of all, the chemical structure of the Myc factor
itself is currently under investigation, and we cannot exclude that several different
molecules contribute to its biological activity, also based on the variety of plant
responses observed. Under this respect, the recent identification, in AM fungal
exudates, of lipochito-oligosaccharides with a structure and effects that are very
similar to those of the rhizobial Nod factors, hints at a further level of crosstalk
between the two symbioses (Maillet et al. 2011). On the other hand, the
upregulation of a steroid-binding protein in response to diffusible AM fungal
signals suggests the involvement of lipid-based signals (Kuhn et al. 2010).
The plant receptors involved in Myc factor perception are also unknown, but the
availability of a few mutant lines affected in AM colonisation has allowed the
identification of genes and proteins that have been positioned along a putative and
only partially characterised signalling pathway.
Signalling and the Re-structuring of Plant Cell Architecture in AM Symbiosis
57
4 The SYM Pathway
Most – although not all – of the responses described so far are in fact mediated by
the activity of a few known gene products (Kistner et al. 2005). These are expressed
in all AM host plants and are known as the “common” SYM genes (Oldroyd and
Downie 2006), with reference to the dual role they have in legumes, in controlling
both AM symbiosis and nitrogen-fixing nodulation (Parniske 2008). While, anyway, a few more genes have been identified in nodulation-specific branches of this
pathway, including the Nod factor receptor-like kinases LjNFR1/5 and MtNFP, no
SYM pathway-related gene has so far been characterised whose activity is unique to
AM interactions.
From a topological perspective (Fig. 2), our partial view of the SYM pathway
shows a very marked directionality from the cell boundaries to the nucleus, which
provides this hypothetical signal transduction pathway with sound rational support.
DMI2 (for Doesn’t Make Infection), the receptor-like kinase of Medicago
truncatula potentially contributing to Myc factor recognition is localised on the
plasma membrane. Three versions of DMI2 homologs are known in angiosperms,
and interestingly the shortest proteins from rice and maize (SYMRK) only recognise
AM fungal signals, whereas the longest homologs, found in legumes, are involved
in the perception of both Myc and Nod factors (Markmann et al. 2008).
All forms expose their kinase domain on the cytoplasmic side of the plasmalemma. The unknown substrate of this kinase is therefore supposed to be a free
cytoplasmic protein. Either this substrate or an indirectly generated signal is
probably reaching the nucleoplasm through nuclear pores. Two nucleoporins,
LjNUP133 (Kanamori et al. 2006) and LjNUP85 (Saito et al. 2007), have in fact
been described in L. japonicus and are indispensable for fungal accommodation and
AM establishment. The fact that such nucleoporins, at least in animals, are not
known to be directly involved in the canonical nuclear pore import/export
mechanisms, leaves anyway the question open about their actual role within the
SYM pathway (Parniske 2008).
Not surprisingly, the nucleus is the organelle where all the remaining SYM gene
products concentrate. The putative cation channels identified as CASTOR and
POLLUX in L. japonicus (Charpentier et al. 2008), homologs of M. truncatula
DMI1 (Ané et al. 2004), are localised on the nuclear envelope and it has been
hypothesized that they could be the cargo of NUP133 and NUP85 (Parniske 2008).
The two last SYM genes involved in AM signalling both localise in the nucleoplasm. MtDMI3 (Lévy et al. 2004) is a kinase whose activity is regulated by the
binding of both calcium and calmodulin through distinct domains. This calcium
dependency gives DMI3 a straightforward role in the downstream transmission of
the signal. Its substrate MtIPD3 (for Interacting Protein of DMI3), homolog of
CYCLOPS in L. japonicus, shows no similarity with proteins of known function,
with the exception of its nuclear localisation signal (Messinese et al. 2007).
This quick glance to the localisation and function of the SYM proteins immediately highlights two central aspects: the identification of the nucleus as the main
58
A. Genre
Fig. 2 The ‘common’ SYM pathway. The panel shows a schematic view of the proteins that are
involved in the so-called SYM pathway, the plant signalling pathway that transduces both signals
from nitrogen-fixing rhizobia (Nod factor) and AM fungi (Myc factor). The first known element of
this common signalling pathway is the receptor-like kinase DMI2 from M. truncatula (homolog
of SYMRK in L. japonicus). Its extracellular domain is thought to be involved in the recognition of
both Nod and Myc factors, while the cytoplasmic domain shows a kinasic activity (lightning sign).
The target of DMI2 is not known, but either this protein or a secondary messenger is supposed to
eventually reach the nucleus. In fact two nucleoporins (NUP85 and NUP133) have been
characterised in L. japonicus, whose functionality is required for downstream signalling. The
nuclear envelope is also the site where DMI1, a putative cation channel homolog of L. japonicus
CASTOR and POLLUX gene products, is localised. The nuclear envelope is a well-known
calcium storage compartment, and the SYM pathway includes the generation of a calcium spiking
signal in the nucleus and perinuclear cytoplasm. Although Ca2+ is not the preferential cargo of
DMI1, a role for this channel has been proposed in the generation of the Ca2+ spiking (Charpentier
et al. 2008). The calcium and calmodulin-dependent kinase DMI3 (CCaMK in L. japonicus) is
localised in the nucleoplasm and is supposed to decode the calcium signal and phosphorylate the
last known protein of the common SYM pathway, IPD3 (homolog of CYCLOPS in L. japonicus),
whose function remains unknown, but localises upstream gene regulation. Please refer to Parniske
(2008) for a compendium of the known SYM gene homologs in different AM host plants
Signalling and the Re-structuring of Plant Cell Architecture in AM Symbiosis
59
target of the signalling pathway and the involvement of calcium as an intermediate
messenger, indispensable for the activation of the nuclear kinase DMI3.
4.1
Calcium as a Secondary Messenger in AM Establishment
While the importance of intranuclear calcium signals was fully appreciated only
after the demonstration of DMI3 nuclear localisation (Smit et al. 2005), the
involvement of Ca2+ signals in AM perception has been postulated since the
identification of a mycorrhizal phenotype in the dmi3 mutants (Lévy et al. 2004).
Sound support to this hypothesis came from the description of calcium spiking in
the perinuclear area (Erhardt et al. 1996; Walker et al. 2000) and more recently in
the nucleoplasm (Sieberer et al. 2009) of root hairs exposed to Nod factors. The first
descriptions of calcium signals in response to AM fungal diffusible factors
(Navazio et al. 2007; Kosuta et al. 2008; Chabaud et al. 2010), although somehow
expected, represented a long awaited support to the theory of the common SYM
pathway.
If, on one hand, the identification of a common signalling pathway controlling
the establishment of two diverse symbioses in legumes has important ecological,
developmental and evolutionary implications (Parniske 2008; Bonfante and Genre
2008; Markmann and Parniske 2009), on the other hand, it raises the problem of
how legumes may distinguish between an AM fungus and a rhizobium and organise
the appropriate accommodation of either one inside the root tissues, when both
signals apparently converge over the same transduction pathway. Under this
respect, stimulating clues come from AM-related calcium signals. Ca2+ is the
most widespread intracellular messenger and transient elevations of its
concentrations couple a wide array of extracellular stimuli to specific physiological
responses (Hetherington and Brownlee 2004; Sanders et al. 2002). With so many
signalling pathways based on the variation of Ca2+ concentration, eukaryotic cells
have evolved sophisticated ways to generate and decode Ca2+ signals in a pathwayspecific manner (the so-called calcium signature). Beside the specific subcellular
localisation of Ca2+ release and the combined action of other intracellular
messengers, plant cells can originate different calcium signatures in response to
different stimuli by changing the duration of the Ca2+ concentration peaks as well as
their frequency (Allen et al. 1999; Harper and Harmon 2005). AM-related nuclear
calcium spikes have in fact been described as less frequent and more irregular
compared to those induced by Nod factors (Sieberer et al. 2009; Chabaud et al.
2010) and recent hypotheses have been proposed, concerning the possible impact of
such frequency modulations in decoding the two respective signals (Kosuta et al.
2008; Hazledine et al. 2009). If this hypothesis will further be confirmed, the
question remains open concerning the mechanism by which the same Ca2+-dependent enzyme, DMI3, can in turn distinguish the two frequency-coded messages.
Due to its positioning within the SYM pathway, calcium spiking can also be used
as a reporter of the activation of this signal transduction pathway. This led, for
60
A. Genre
example, to the identification of AM-responsive districts in the root epidermis,
localised between 1 and 2 cm from the tip of young lateral roots (Chabaud et al.
2010). Further studies are expected to clarify to what extent the calcium spiking
response (and hence the SYM pathway) is also involved after cell entry, for fungal
colonization of epidermal and cortical cells. Under this respect, the observation of
intense nuclear calcium spiking in the small group of epidermal cells immediately
surrounding each AM hyphopodium (Chabaud et al. 2010) confirmed that the
response is directly related to the colonization process and that the SYM pathway
is active in epidermal cells. This was previously only suggested by the phenotype of
SYM gene mutants where fungal growth was arrested within the epidermal layer
(Kistner et al. 2005; Novero et al. 2002) and gene expression was altered in the
same cells (Chabaud et al. 2002).
5 Fungal Accommodation
Upon the development of a hyphopodium on the outer wall of root epidermal cells
both the plant and fungus enter a novel developmental program that culminates in
fungal accommodation inside the root cell lumen and symbiosis full functionality.
Among the epidermal cells where the nuclear calcium spiking is observed, a few of
those in direct contact with the hyphopodium dramatically reorganise their cytoplasm. The first visible sign of this response is the migration of the nucleus at the
fungus contact site (Genre et al. 2005). Interestingly, such repositioned nuclei show
the highest spiking frequencies (Chabaud et al. 2010), suggesting that the initiation
of the accommodation program correlates with a strong activation of the SYM
pathway – whose nucleus-oriented directionality has been discussed above. During
the following few hours, the repositioned nucleus is progressively surrounded by an
aggregation of cytoplasm, which expands into a columnar shape as the nucleus
moves again from the contact site to the opposite side of the cell. Such a broad and
roughly linear cytoplasmic aggregation is called the prepenetration apparatus (or
PPA; Genre et al. 2005), as it predicts the intracellular path where the penetration
hypha will grow. The PPA includes cytoskeletal fibres as well as the whole
exocytotic apparatus, from a large accumulation of endoplasmic reticulum to
numerous Golgi stacks, trans-golgi network and secretory vesicles (Genre et al.
2005, 2008). Remarkably, PPA assembly requires 6–7 h after hyphopodium adhesion. During this time, hyphopodium development appears to be totally arrested,
which strongly suggests that fungal growth, at least at this stage, is controlled by the
plant process that prepares cell entry.
The direct cell-to-cell contact may favour an intense and rapid signal exchange,
which could include – but not be limited to – diffusible compounds such as the Myc
factor. Based on the comparison with plant pathogenic interactions, the release of
effector molecules from the hyphopodium seems likely (Catanzariti et al. 2006;
O’Connell and Panstruga 2006; Valent and Khang 2010). Indeed, the observed
interpenetration of fungal and plant cell walls at the contact site (Bonfante and
Signalling and the Re-structuring of Plant Cell Architecture in AM Symbiosis
61
Genre 2010) suggests an intense local remodelling of the apoplast, which is
compatible with protein secretion and bioactive molecules release (O’Connell
and Panstruga 2006).
On the plant side, little information is available concerning secreted molecules
that may be active during the initiation of the symbiotic phase. Most of the host
plant known mutants showing a block to epidermal cell penetration by the fungus
bear mutations in one of the SYM gene homologs (Parniske 2008). Nevertheless,
the screening of mutagenised lines from tomato, maize and Petunia has led to the
identification of a few mutants whose phenotype involves the inhibition of fungal
proliferation, including abnormal hyphopodium development (Barker et al. 1998;
David-Schwartz et al. 2001; Paszkowski et al. 2006; Reddy et al. 2007). At least in
tomato, such effects are cancelled by the presence of WT plants in the vicinity of
the mutants, strongly suggesting that the inhibition of fungal proliferation is due to
the lack of so-far unidentified root secreted molecules (Cavagnaro et al. 2004).
The occurrence of signalling events during AM colonisation of the inner root
tissues is suggested by the observation of long-distance responses in the plant cells
(Fig. 3). As soon as the PPA is being completed in an epidermal cell, nuclear
repositioning in front of its terminal site is reported in the underlying cortical cell,
associated with the initiation of a new PPA (Genre et al. 2008). This implies that the
site where the fungus will cross the epidermal/cortical cell wall is being marked
before the fungus has even started penetrating the epidermal cell. Analogous and
even more striking clues for the involvement of long-range signals are observed in
the inner cortex of Paris-type mycorrhizas, such as those that develop in carrot.
Here, fungal colonisation of the root tissue proceeds from cell to cell. PPAs in
progressively advanced stages of development are observed in several cells ahead
of the developing hyphal tip, indicating that a signal triggering the accommodation
process is translocated on a relatively long distance (Genre et al. 2008). Lastly,
plant cell responses in the non-colonised cells adjacent to arbuscules have been
detected with both cellular and molecular methods. Microtubule rearrangement in
the vicinity of arbusculated cells of M. truncatula (Blancaflor et al. 2001) and gene
regulation in non-colonized cortical cells from the colonised areas of M. truncatula
(Harrison 2005) and tomato roots (Balestrini et al. 2007) have both been interpreted
as the result of cell-to-cell signal transfer. Interestingly, lysophosphatidylcholine
extracted from arbuscule-containing roots has been shown to induce the
upregulation of AM-specific phosphate transporters in non-colonised roots,
suggesting that this molecule could be one of such cell-to-cell signals (Drissner
et al. 2007).
Intense membrane dynamics occur within the PPA, as documented by the rapid
uptake of the endocytotic marker FM4-64 (Genre et al. 2005) and the simultaneous
presence of exocytotic vesicles and endosomes (Genre et al. 2008). Endosomemediated signalling is a well-known mechanism, which, especially in plant cells,
grants the rapid translocation of active receptors from the perception site on the
plasma membrane to the cellular domain where the response is started (Geldner and
Robatzek 2008). From this point of view one can reasonably speculate that the PPA
acts as a route for signalling endosomes, thanks to the dynamism of its vesicular
62
A. Genre
Fig. 3 Long-range signal transmission during AM colonisation of the root tissues. Several cellular
clues indicate the existence of diffusible signals that induce prepenetration responses in the inner
root tissues before the cells are in direct contact with the fungus. The diagram in (a) shows the
initiation of a PPA in the outer cortical cell, in line with the fully assembled PPA of the overlaying
epidermal cell. The diagram in (b) shows a similar but more extensive coordination in PPA
assembly within a group of inner cortical cells. Prepenetration responses appear to be progressively less advanced with increasing distance from the growing hyphal tip, but PPA alignment
along the cell file is evident. The diagram in (c) summarises the observation of arbuscule-specific
gene regulation and cytoskeleton remodelling in non-colonised cells adjacent to the arbusculated
ones. Both are suggestive of a signal diffusion from the arbuscule to activate the neighbouring cells
in preparation for potential fungal entry
content. The longitudinal orientation of microtubules and microfilaments inside the
PPA aggregate (Genre et al. 2005) also supports this view. This would explain how
prepenetration responses coordinate between adjacent cells, but does not clarify
whether the upstream stimulus has a direct fungal origin or derives from a secondary plant signal. The location of such responses deep within the root cells and
tissues currently hinders our attempts to solve this conundrum. Only the availability
of genomic information and the application of genetic tools on the fungus will
likely shed light on such aspects.
A remarkable trait stands out from the study of prepenetration responses in AM
host plants: nuclear movement, cytoskeleton arrangement, focal concentration of
exocytotic elements and the apparent directional translocation of signalling factors
are suggestive of cell polarisation processes. The onset of polarity is common to many
plant developmental processes, including responses to pathogens (Genre et al. 2009)
Signalling and the Re-structuring of Plant Cell Architecture in AM Symbiosis
63
but also apical growth and hormonal fluxes. And indeed the directional transport of
auxin is known to be involved in the establishment of cell polarity in a self-inducing
mechanism (Lee and Cho 2006; Feraru and Friml 2008).
Nevertheless, a correlation between AM colonisation and hormone activity has
so far only been highlighted on organ and systemic scale. Auxin levels increase in
mycorrhizal root systems, but are unrelated to the distribution of colonised areas
(Jentschel et al. 2007). Auxin-upregulated genes, such as the leghemoglobin Lb29
from Vicia faba, were shown to be also induced during AM colonization, indirectly
confirming the AM-induced increase in auxin levels (Ludwig-M€uller and G€uther
2007). The effect of AM on ethylene, abscisic acid and jasmonate-related
compounds, as well as the expression of key genes in hormone biosynthetic
pathways shows important differences depending on the fungal species (LópezRáez et al. 2010). Even more controversial is the involvement of jasmonic and
salicylic acids in regulating AM-induced defense (Gutjahr and Paszkowski 2009;
Hause and Schaarschmidt 2009). All these data indicate a role for hormone-based
signals especially on the systemic effects of the AM interaction, which range from
the long described “growth effect” observed in mycorrhizal plants (Smith and Read
2008) to the large-scale reprogramming of regulatory networks as it is being
highlighted by transcriptomic analyses and microarray assays (Liu et al. 2007;
Fiorilli et al. 2009). Furthermore, beside plant hormones, long distance signals
might also include mobile microRNAs, as recently suggested by expression
analyses (Gu et al. 2010; Branscheid et al. 2010).
6 Signalling During Symbiosis Functioning
Arbuscule appearance inside the lumen of inner cortical cells hallmarks the establishment of mutualism. The extensive periarbuscular interface and membrane in
fact mediate the intense nutrient exchange that characterises AM symbiosis and
represents the basis of its ecological and evolutionary success. Signalling
mechanisms partly similar to those controlling intracellular fungal growth in the
outer tissues are predicted to control arbuscule development also, as suggested by
the reiteration of the PPA model prior to the development of new fungal branches
(Genre et al. 2008). Nonetheless, the accommodation program within the inner
cortex develops unique processes, which can be envisaged through both cellular
and molecular investigations. The genetic control of arbuscule development is only
partly ruled by the SYM pathway. Most SYM gene mutants either block fungal
penetration in the outer cell layers or develop normal and functional arbuscules
when such surface block is taken over (Morandi et al. 2005). Very few exceptions
are known: a castor mutant (LjSym4-1), in L. japonicus, was shown to develop
fewer arbuscules and more intercellular hyphae than the wild type, suggesting that
the mutation affects cortical cell entry (Novero et al. 2002). A very similar
phenotype derives from RNAi-mediated downregulation of the Vapyrin gene in
M. truncatula, but the gene does not belong to the SYM group (Pumplin et al.
64
A. Genre
2010). Interestingly, the Vapyrin gene is unique to the plant kingdom but absent in
the AM non-host Arabidopsis, suggesting a possible AM-specific – although
unknown – function for the encoded ankyrin-like protein.
Morphological evidence for the diversity of the cell responses in inner cortical
versus epidermal and outer cortical cells include the overall morphology of the
PPAs (thinner and roughly linear in the outer tissues, broader and bulged in the
inner cortex), the final positioning of the nucleus (which persists in the cell centre
during the whole arbuscule life span), the decondensation of chromatin, only
observed in the inner cortex and suggestive of more intense transcriptional activity
(Genre et al. 2008). How all these processes are controlled and specifically triggered in the cells that will contain arbuscules remains to be clarified.
When arbuscules reach maturity, filling up most of the cell volume, the nutrients
they transfer also seem to have a role as signals. Javot et al. (2007) have elegantly
demonstrated that if the plant does not receive P from the fungus – due to either
RNAi silencing or mutation of the arbuscule-specific MtPT4 phosphate transporter –
cortical cell colonisation is followed by an anticipated process of fungal senescence,
resulting in small arbuscules made of thick, short branches. Such arbuscules septate
and die much earlier (3–4 days after inoculation) than normal arbuscules developed
in wild-type plants (with a reported life span of over 8 days). These observations
indicate that phosphate, the most important nutrient provided to host plants by AM
fungi, can act upstream a signalling pathway that controls arbuscule morphogenesis
and senescence. Although this result was obtained by knocking down a plant
transporter, it strongly suggests that the fungal efficiency in P delivery has a
major importance for symbiosis establishment.
Altogether, the occurrence of signal exchanges inside the arbusculated cells is
more than just likely. A strict coordination between the two partners is required
during the whole development of the interaction, and the complexity of arbuscule
differentiation and functioning makes these structures the site where plant–fungus
signalling must inevitably reach the highest intensity. Among such signals, the very
nutrients that the plant and the fungus exchange through the periarbuscular interface are to be considered as an important and perhaps major component. It is
expected that over the next few years AM-related research, currently focussed on
the presymbiotic signals, will concentrate on the signalling events that associate
with arbuscule development, function and senescence.
7 Conclusions
AM symbiosis is the result of a delicate balance, where both partners release
signals, respond to each other and coordinate their developmental programs, leading to the accommodation of a eukaryote inside another eukaryote. This rather
uncommon event presents unique peculiarities in the case of AM. The hosted
organism, in fact, as a true endosymbiont, develops inside the plant cells; nevertheless, its intracellular structures maintain direct communication with the extraradical
Signalling and the Re-structuring of Plant Cell Architecture in AM Symbiosis
65
mycelium that explores the soil. The absence of septa and the strong cytoplasmic
streams that characterise AM fungi lead to a condition where a true cellular
continuity can be envisaged within the whole organism.
Compared to such a complex biological system, our current view of the cellular
and molecular mechanisms that rule AM is not simply limited, but often biased.
First, a substantial lack of information concerning the fungus impacts both our
knowledge and the overall picture that we get of AM interactions. Besides leaving
many fundamental aspects of AM fungal biology completely obscure, such a ‘green
shift’ is responsible for the common idea that AM host plants hold major control
over the symbiosis. This is supported by many experiments, where the mutation of a
single plant gene results in the block of fungal development at the hyphopodium
stage. Nevertheless, the fact that fungal mutants have not so far been obtained does
not exclude that fungal control is just as strong. Similarly, plant genetic manipulation has allowed detailed investigations of the cell responses during fungal
colonisation, but we know very little about the morphogenetical programs that
originate all of the symbiotic structures of AM fungi, from hyphopodia to intracellular hyphae, coils, vesicles and arbuscules. Once again, the lack of appropriate
tools strongly affects our possibilities to investigate these aspects. The achievement
of transient genetic transformation in Glomus intraradices (Helber and Requena
2008) envisages the dawn of a new era in AM research, where this methodological
gap can be filled.
A somehow similar bias can be ascribed to the appeal of simplification. The
advancements that have been made on the identification of the SYM signalling
pathway, far from providing a complete signal transduction sequence, have
polarised the attention of researchers on this one signalling mechanism. It should
anyway be noticed that an increasing set of data demonstrate the existence of signal
transduction processes in AM that transcend the SYM pathway. Recent examples
include the regulation of AM-specific marker genes in SYM mutants (Gutjahr et al.
2008; Kuhn et al. 2010). Besides being still partially obscure, the presymbiotic
signalling in AM may thus be significantly more complex than expected and
research in the next years is likely to provide us with both answers and new
questions.
Altogether, the signals and responses that we know to be controlling AM
establishment fall into well-known mechanisms of all eukaryotic cells. Signal
transduction through elicitor perception, receptor activation, calcium-mediated
signals, phosphorylation cascades; cell responses involving cytoplasm aggregation
and focal exocytosis; and the reprogramming of metabolic and physiological
pathways on a local and systemic scale. All of these events are common to most
plant (and animal) interactions. A challenge for the near future will be to understand
how the same basic mechanisms are routed towards one conclusion or another. For
example, cell responses that involve nuclear repositioning, cytoplasmic aggregation, cytoskeletal remodelling and local membrane dynamics are described in both
pathogenic and symbiotic interactions, but the same cellular machinery is obviously
able to organise defense responses, such as the development of cell wall appositions
underneath the penetration peg of a pathogen, or accommodation processes, such as
66
A. Genre
the assembly of the perifungal interface to host an AM fungus. Molecular
mechanisms such as the regulation of defense-related gene expression are also
known to be involved in both mutualistic and pathogenic interactions (Paszkowski
2006b), but the control, modulation and reprogramming of such responses must
depend on specific signals and signalling cascades that identify each interacting
microorganism.
If, on one hand, presymbiotic signals have been one of the most difficult
conundrums that AM research has faced so far, on the other hand, they take place
during the most accessible phase of AM development. The plant and fungus are still
separated and can be manipulated independently to highlight the effects of diffusible molecules and study the details of cellular and molecular responses. But
starting from hyphopodium adhesion, the tools we can apply to address the investigation of signalling events inside the root tissues are indeed less powerful. Many
questions remain therefore open. For example, experimental evidence shows that
the PPA predicts intracellular fungal growth, but how is the penetrating hyphal tip
induced to exactly follow that membranous track, rather than break into the plant
cytoplasm? Similarly, the prepenetration responses that develop in the cortical cells
do not extend indefinitely. Each infection event leads to the colonisation of a
limited zone of the root, but how such limits are set and how fungal colonisation
of the cortex is controlled by either the plant or the fungus remains largely
unexplored. Such examples strongly suggest the existence of a complex network
of signalling mechanisms throughout the AM symbiotic cycle, and these clues will
likely be the germ of AM signal research for the next years.
Acknowledgements I am grateful to Paola Bonfante and Luisa Lanfranco for useful discussion
and critical reading of the manuscript. Experimental work described here was supported by grants
to Paola Bonfante from Compagnia di San Paolo and Converging technologies-CIPE BIOBIT
project.
References
Akiyama K, Matsuzaki K, Hayashi H (2005) Plant sesquiterpenes induce hyphal branching in
arbuscular mycorrhizal fungi. Nature 435:824–827
Akiyama K, Ogasawara S, Ito S, Hayashi H (2010) Structural requirements of strigolactones for
hyphal branching in AM fungi. Plant Cell Physiol 51:1104–1117
Allen GJ, Kwak JM, Chu SP, Llopis J, Tsien RY, Harper JF, Schroeder JI (1999) Cameleon
calcium indicator reports cytoplasmic calcium dynamics in Arabidopsis guard cells. Plant J
19:735–747
Ané JM, Kiss GB, Riely BK, Penmetsa RV, Oldroyd GED, Ayax C, Lévy J, Debellé F, Baek J-M,
Kalo P, Rosenberg C, Roe BA, Long SR, Dénarie J, Cook DR (2004) Medicago truncatula
DMI1 required for bacterial and fungal symbioses in legumes. Science 303:1364–1367
Arite T, Umehara M, Ishikawa S, Hanada A, Maekawa M, Yamaguchi S, Kyozuka J (2009) d14, a
strigolactone insensitive mutant of rice, shows an accelerated outgrowth of tillers. Plant Cell
Physiol 50:1416–1424
Balestrini R, Gómez-Ariza J, Lanfranco L, Bonfante P (2007) Laser microdissection reveals that
transcripts for five plant and one fungal phosphate transporter genes are contemporaneously
present in arbusculated cells. Mol Plant -Microbe Interact 20:1055–1062
Signalling and the Re-structuring of Plant Cell Architecture in AM Symbiosis
67
Barker SJ, Stummer B, Gao L, Dispain I, O’Connor PJ, Smith SE (1998) A mutant in Lycopersicon
esculentum Mill, with highly reduced VA mycorrhizal colonization: isolation and preliminary
characterization. Plant J 15:791–797
Bastmeyer M, Deising H, Bechinger C (2002) Force exertion in fungal infection. Annu Rev
Biophys Biomol Struct 31:321–341
Bécard G, Fortin JA (1988) Early events of vesicular-arbuscular mycorrhiza formation on Ri TDNA transformed roots. New Phytol 108:211–218
Besserer A, Puech-Pagès V (2006) Strigolactones stimulate arbuscular mycorrhizal fungi by
activating mitochondria. PLoS Biol 4:1239–1247
Besserer A, Becard G, Jauneau A, Roux C, Sejalon-Delmas N (2008) GR24, a synthetic analog of
strigolactones, stimulates the mitosis and growth of the arbuscular mycorrhizal fungus
Gigaspora rosea by boosting its energy metabolism. Plant Physiol 148:402–413
Blancaflor EB, Zhao L, Harrison MJ (2001) Microtubule organization in root cells of Medicago
truncatula during development of an arbuscular mycorrhizal symbiosis with Glomus
versiforme. Protoplasma 217:154–165
Bonfante P (2001) At the interface between mycorrhizal fungi and plants: the structural organization of cell wall, plasma membrane and cytoskeleton. In: Hock B (ed) Mycota, vol IX, Fungal
associations. Springer, Heidelberg, pp 45–91
Bonfante P, Genre A (2008) Plants and arbuscular mycorrhizal fungi: an evolutionary-developmental
perspective. Trends Plant Sci 13:492–498
Bonfante P, Genre A (2010) Mechanisms underlying beneficial plant–fungus interactions in
mycorrhizal symbiosis. Nature Commun 1:48
Branscheid A, Sieh D, Pant BD, May P, Devers EA, Elkrog A, Schauser L, Scheible WR, Krajinski F
(2010) Expression pattern suggests a role of MiR399 in the regulation of the cellular response to
local Pi increase during arbuscular mycorrhizal symbiosis. Mol Plant Microbe Interact
23:915–926
Buee M, Rossignol M, Jauneau A, Ranjeva R, Becard G (2000) The pre-symbiotic growth of
arbuscular mycorrhizal fungi is induced by a branching factor partially purified from plant root
exudates. Mol Plant Microbe Interact 13:693–698
Cano C, Dickson S, González-Guerrero M, Bago A (2008) In vitro cultures open new prospects for
basic research in arbuscular mycorrhizas. In: Mycorrhiza: state of the art, genetics and
molecular biology, eco-function, biotechnology, eco-physiology, structure and systematics,
3 rd ed. Springer, Heidelberg, pp 627–654
Catanzariti AM, Dodds PN, Lawrence GJ, Ayliffe MA, Ellisa JG (2006) Haustorially expressed
secreted proteins from flax rust are highly enriched for avirulence elicitors. Plant Cell
18:243–256
Cavagnaro TR, Smith FA, Hay G, Carne-Cavagnaro VL, Smith SE (2004) Inoculum type does not
affect overall resistance of an arbuscular mycorrhiza-defective tomato mutant to colonisation
but inoculation does change competitive interactions with wild-type tomato. New Phytol
161:485–494
Chabaud M, Venard C, Defaux-Petras A, Becard G, Barker DG (2002) Targeted inoculation of
Medicago truncatula in vitro root cultures reveales MtENOD11 expression during early stages
of infection by arbuscular mycorrhizal fungi. New Phytol 156:265–273
Chabaud M, Genre A, Sieberer BJ, Faccio A, Fournier J, Novero M, Barker DG, Bonfante P (2010)
Arbuscular mycorrhizal hyphopodia and germinated spore exudates trigger Ca2+ spiking in the
legume and nonlegume root epidermis. New Phytol 189:347–355
Charpentier M, Bredemeier R, Wanner G, Takeda N, Schleiff E, Parniske P (2008) Lotus
japonicus CASTOR and POLLUX are ion channels essential for perinuclear calcium spiking
in legume root endosymbiosis. Plant Cell 20:3467–3479
Cook CE, Whichard LP et al (1966) Germination of witchweed (Striga Lutea Lour): isolation and
properties of a potent stimulant. Science 154:1189–1190
Croll D, Giovannetti M, Koch A, Sbrana C, Ehinger M, Lammers P, Sanders I (2009) Nonself
vegetative fusion and genetic exchange in the arbuscular mycorrhizal fungus Glomus
intraradices. New Phytol 181:924–937
68
A. Genre
David-Schwartz R, Badani H, Smadar W, Levy AA, Galili G, Kapulnik Y (2001) Identification of
a novel genetically controlled step in mycorrhizal colonization: plant resistance to infection by
fungal spores but not extra-radical hyphae. Plant J 27:561–569
Delaux PM, Bécard G, Séjalon-Delmas N (2009) Evolutive origin of strigolactones. Proceedings
of the 6th international conference on Mycorrhiza, 9–14 August 2009. International Mycorrhiza Society, Belo Horizonte, Brazil
Dickson S (2004) The Arum–Paris continuum of mycorrhizal symbioses. New Phytol
163:187–200
Dickson S, Kolesik P (1999) Visualisation of mycorrhizal fungal structures and quantification of
their surface area and volume using laser scanning confocal microscopy. Mycorrhiza
9:205–213
Drissner D, Kunze G, Callewaert N, Gehrig P, Tamasloukht MB, Boller T, Felix G, Amrhein N,
Bucher M (2007) Lysophosphatidylcholine is a signal in the arbuscular mycorrhizal symbiosis.
Science 318:265–268
Erhardt DW, Wais R, Long SR (1996) Calcium spiking in plant root hairs responding to Rhizobium
nodulation signals. Cell 85:673–681
Feraru E, Friml J (2008) PIN polar targeting. Plant Physiol 147:1553–1559
Fiorilli V, Catoni M, Miozzi L, Novero M, Accotto GP, Lanfranco L (2009) Global and cell-type
gene expression profiles in tomato plants colonized by an arbuscular mycorrhizal fungus. New
Phytol 184:975–987
Geldner N, Robatzek S (2008) Plant receptors go endosomal: a moving view on signal transduction. Plant Physiol 147:1565–1574
Genre A, Chabaud M, Timmers T, Bonfante P, Barker DG (2005) Arbuscular mycorrhizal fungi
elicit a novel intracellular apparatus in Medicago truncatula root epidermal cells before
infection. Plant Cell 17:3489–3499
Genre A, Chabaud M, Faccio A, Barker DG, Bonfante P (2008) Prepenetration apparatus assembly
precedes and predicts the colonization patterns of arbuscular mycorrhizal fungi within the root
cortex of both Medicago truncatula and Daucus carota. Plant Cell 20:1407–1420
Genre A, Ortu G, Bertoldo C, Martino E, Bonfante P (2009) Biotic and abiotic stimulation of root
epidermal cells reveals common and specific responses to arbuscular mycorrhizal fungi. Plant
Physiol 149:1424–1434
Gómez-Roldán V, Fermas S, Brewer PB, Puech-Pagès V, Dun EA, Pillot JP, Letisse F, Matusova R,
Danoun S, Portais JC, Bouwmeester H, Bécard G, Beveridge CA, Rameau C, Rochange SF
(2008) Strigolactone inhibition of shoot branching. Nature 455:189–194
Gu M, Xu K, Chen A, Zhu Y, Tang G, Xu G (2010) Expression analysis suggests potential roles of
microRNAs for phosphate and arbuscular mycorrhizal signaling in Solanum lycopersicum.
Physiol Plant 138:226–237
Guether M, Neuhauser B, Balestrini R, Dynowski M, Ludewig U, Bonfante P (2009) A mycorrhizal specific ammonium transporter from Lotus japonicus acquires nitrogen. Plant Physiol
150:73–83
Gutjahr C, Paszkowski U (2009) Weights in the balance: jasmonic acid and salicylic acid signaling
in root-biotroph interactions. Mol Plant Microbe Interact 22:763–772
Gutjahr C, Banba M, Croset V, An K, Miyao A, An G, Hirochika H, Imaizumi-Anraku H,
Paszkowski U (2008) Arbuscular mycorrhiza-specific signaling in rice transcends the common
symbiosis signaling pathway. Plant Cell 20:2989–3005
Gutjahr C, Novero M, Guether M, Montanari O, Udvardi M, Bonfante P (2009) Presymbiotic
factors released by the arbuscular mycorrhizal fungus Gigaspora margarita induce starch
accumulation in Lotus japonicus roots. New Phytol 183:53–61
Harper JF, Harmon A (2005) Plants, symbiosis and parasites: a calcium signalling connection. Nat
Rev Mol Cell Biol 6:555–566
Harrison MJ (2005) Signaling in the arbuscolar mycorrhizal symbiosis. Annu Rev Microbiol
59:19–42
Hause B, Schaarschmidt S (2009) The role of jasmonates in mutualistic symbioses between plants
and soil-born microoganisms. Phytochemistry 70:1589–1599
Signalling and the Re-structuring of Plant Cell Architecture in AM Symbiosis
69
Hazledine S, Sun J, Wysham D, Downie JA, Oldroyd GE, Morris RJ (2009) Nonlinear time series
analysis of nodulation factor induced calcium oscillations: evidence for deterministic chaos?
PLoS One 4:e6637
Helber N, Requena N (2008) Expression of the fluorescence markers DsRed and GFP fused to a
nuclear localization signal in the arbuscular mycorrhizal fungus Glomus intraradices. New
Phytol 177:537–548
Hetherington AM, Brownlee C (2004) The generation of Ca2+ signals in plants. Annu Rev Plant
Biol 55:401–427
Hijri M, Sanders IR (2005) Low gene copy number shows that arbuscular mycorrhizal fungi
inherit genetically different nuclei. Nature 433:161–163
Ishikawa S, Maekawa M, Arite T, Onishi K, Takamure I, Kyozuka J (2005) Suppression of tiller
bud activity in tillering dwarf mutants of rice. Plant Cell Physiol 46:79–86
Javot H, Penmetsa RV, Terzaghi N, Cook DR, Harrison MJ (2007) A Medicago truncatula
phosphate transporter indispensable for the arbuscular mycorrhizal symbiosis. Proc Natl
Acad Sci USA 104:1720–1725
Jentschel K, Thiel D, Rehn F, Ludwig-M€
uller J (2007) Arbuscular mycorrhiza enhances auxin
levels and alters auxin biosynthesis in Tropaeolum majus during early stages of colonization.
Physiol Plant 129:320–333
Kanamori N, Madsen LH, Radutoiu S, Frantescu M, Quistgaard EMH, Miwa H, Downie JA,
James EK, Felle HH, Haaning LL (2006) A nucleoporin is required for induction of Ca2+
spiking in legume nodule development and essential for rhizobial and fungal symbiosis. Proc
Natl Acad Sci USA 103:359–364
Kistner C, Winzer T, Pitzschke A, Mulder L, Sato S, Kaneko T, Tabata S, Sandal N, Stougaard J,
Webb KJ, Szczyglowski K, Parniske M (2005) Seven Lotus japonicus genes required for
transcriptional reprogramming of the root during fungal and bacterial symbiosis. Plant Cell
17:2217–2229
Kobae K, Hata S (2010) Dynamics of periarbuscular membranes visualized with a fluorescent
phosphate transporter in arbuscular mycorrhizal roots of rice. Plant Cell Physiol 51:341–353
Koltai H, LekKala SP, Bahattacharya C, Mayzlish-Gati E, Resnick N, Wininger S, Dor E,
Yoneyama K, Yoneyama K, Hershenhorn J, Joel DM, Kapulnik YC (2010) A tomato
strigolactone-impaired mutant displays aberrant shoot morphology and plant interactions.
J Exp Bot 61:1739–1749
Kosuta S, Chabaud M, Lougnon G, Gough C, Dénarié J, Barker DG, Bécard G (2003) A diffusible
factor from arbuscular mycorrhizal fungi induces symbiosis-specific MtENOD11 expression in
roots of Medicago truncatula. Plant Physiol 131:952–962
Kosuta S, Hazledine S, Sun J, Miwa H, Morris RJ, Downie JA, Oldroyd GE (2008) Differential
and chaotic calcium signatures in the symbiosis signaling pathway of legumes. Proc Natl Acad
Sci USA 105:9823–9828
Kuhn H, Kuster H, Requena N (2010) Membrane steroid-binding protein 1 induced by a diffusible
fungal signal is critical for mycorrhization in Medicago truncatula. New Phytol 185:716–733
Lee SH, Cho HT (2006) PINOID positively regulates auxin efflux in Arabidopsis root hair cells
and tobacco cells. Plant Cell 18:1604–1616
Leeper GW (1952) Factors affecting availability of inorganic nutrients in soils with special
reference to micronutrient metals. Annu Rev Plant Physiol 3:1–16
Leight J, Hodge A, Fitter AH (2009) Arbuscular mycorrhizal fungi can transfer substantial
amounts of nitrogen to their host plant from organic material. New Phytol 181:199–207
Lévy J, Bres C, Geurts R, Chalhoub B, Kulikova O, Duc G, Journet EP, Ané JM, Laubert E,
Bisseling T, Dénarié J, Rosenberg C, Debellé F (2004) A putative Ca2+ and calmodulindependent protein kinase required for bacterial and fungal symbioses. Science 303:1361–1364
Liu JY, Maldonado-Mendoza I, Lopez-Meyer M, Cheung F, Town CD, Harrison MJ (2007)
Arbuscular mycorrhizal symbiosis is accompanied by local and systemic alterations in gene
expression and an increase in disease resistance in the shoots. Plant J 50:529–544
70
A. Genre
López-Ráez JA, Verhage A, Fernández I, Garcı́a JM, Azcón-Aguilar C, Flors V, Pozo MJ (2010)
Hormonal and transcriptional profiles highlight common and differential host responses to
arbuscular mycorrhizal fungi and the regulation of the oxylipin pathway. J Exp Bot
61:2589–2601
Ludwig-M€uller J, G€uther M (2007) Auxins as signals in arbuscular mycorrhiza formation. Plant
Signal Behav 2:194–196
Maillet F, Poinsot V, Andre O, Puech-Pages V, Haouy A, Gueunier M, Giraudet D, Formey D,
Martinez EA, Driguez H, Becard G, Dénarié J (2011) Fungal lipochitooligosaccharide symbiotic signals in arbuscular mycorrhiza. Nature 69:58–63
Markmann K, Parniske M (2009) Evolution of root endosymbiosis with bacteria: how novel are
nodules? Trends Plant Sci 14:77–86
Markmann K, Giczey G, Parniske M (2008) Functional adaptation of a plant receptor-kinase paved
the way for the evolution of intracellular root symbioses with bacteria. PLoS Biol 6:e68
Messinese E, Mun JH, Yeun LH, Jayaraman D, Rougé P, Barre A, Lougnon G, Schornack S, Bono JJ,
Cook DR, Ané JM (2007) A novel nuclear protein interacts with the symbiotic DMI3 calciumand calmodulin-dependent protein kinase of Medicago truncatula. Mol Plant Microbe Interact
20:912–921
Morandi D, Prado E, Sagan M, Duc G (2005) Characterisation of new symbiotic Medicago
truncatula (Gaertn.) mutants, and phenotypic or genotypic complementary information on
previously described mutants. Mycorrhiza 15:283–289
Navazio L, Moscatiello R, Genre A, Novero M, Baldan B, Bonfante P, Mariani P (2007) A
diffusible signal from arbuscular mycorrhizal fungi elicits a transient cytosolic calcium elevation in host plant cells. Plant Physiol 144:673–681
Novero M, Faccio A, Genre A, Stougaard J, Webb K, Mulder L, Parniske M, Bonfante P (2002)
Dual requierment of the LjSYM4 gene for mycorrhizal development in epidermal and cortical
cells of Lotus japonicus roots. New Phytol 154:741–749
O’Connell RJ, Panstruga R (2006) Tête à tête inside a plant cell: establishing compatibility
between plants and biotrophic fungi and oomycetes. New Phytol 171:699–718
Olah B, Briere C, Becard G, Denarie J, Gough C (2005) Nod factors and a diffusible factor from
arbuscular mycorrhizal fungi stimulate lateral root formation in Medicago truncatula via the
DMI1/DMI2 signalling pathway. Plant J 44:195–207
Oldroyd GED, Downie JA (2006) Nuclear calcium changes at the core of symbiosis signalling.
Curr Opin Plant Biol 9:351–357
Parniske M (2008) Arbuscular mycorrhiza: the mother of plant root endosymbioses. Nat Rev
Microbiol 6:763–775
Paszkowski U (2006a) A journey through signaling in arbuscular mycorrhizal symbioses. New
Phytol 172:35–46
Paszkowski U (2006b) Mutualism and parasitism: the yin and yang of plant symbioses. Curr Opin
Plant Biol 9:364–370
Paszkowski U, Jakovleva L, Boller T (2006) Maize mutants affected at distinct stages of the
arbuscular mycorrhizal symbiosis. Plant J 47:165–173
Pawlowska TE, Taylor JW (2004) Organization of genetic variation in individuals of arbuscular
mycorrhizal fungi. Nature 427:733–737
Pumplin N, Harrison MJ (2009) Live-cell imaging reveals periarbuscular membrane domains and
organelle location in Medicago truncatula roots during arbuscular mycorrhizal symbiosis.
Plant Physiol 151:809–819
Pumplin N, Mondo SJ, Topp S, Starker CG, Gantt JS, Harrison MJ (2010) Medicago truncatula
Vapyrin is a novel protein required for arbuscular mycorrhizal symbiosis. Plant J 61:482–494
Reddy DMRS, Schorderet M, Feller U, Reinhardt D (2007) A petunia mutant affected in intracellular accommodation and morphogenesis of arbuscular mycorrhizal fungi. Plant J 51:739–750
Remy W, Taylor TN, Hass H, Kerp H (1994) Four hundred-million-year-old vesicular arbuscular
mycorrhizae. Proc Natl Acad Sci USA 91:11841–11843
Signalling and the Re-structuring of Plant Cell Architecture in AM Symbiosis
71
Saito K, Yoshikawa M, Yano K, Miwa H, Uchida H, Asamizu E, Sato S, Tabata S, ImaizumiAnraku H, Umehara Y, Kouchi H, Murooka Y, Szczyglowski K, Downie JA, Parniske M,
Makoto Hayashi M, Masayoshi Kawaguchi M (2007) NUCLEOPORIN85 is required for
calcium spiking, fungal and bacterial symbioses, and seed production in Lotus japonicus.
Plant Cell 19:610–624
Sanders IR, Croll D (2010) Arbuscular mycorrhiza: the challenge to understand the genetics of the
fungal partner. Annu Rev Genet 44:271–292
Sanders D, Pelloux J, Brownlee C, Harper JF (2002) Calcium at the crossroads of signaling. Plant
Cell 14:S401–S417
Sbrana C, Giovannetti M (2005) Chemotropism in the arbuscular mycorrhizal fungus Glomus
mosseae. Mycorrhiza 15:539–545
Sieberer BJ, Chabaud M, Timmers AC, Monin A, Fournier J, Barker DG (2009) A nucleartargeted cameleon demonstrates intranuclear Ca2+ spiking in Medicago truncatula root hairs in
response to rhizobial nodulation factors. Plant Physiol 151:1197–1206
Smit P, Raedts J, Portyanko V, Debellé F, Gough C, Bisseling T, Geurts R (2005) NSP1 of the
GRAS protein family is essential for rhizobial Nod factor-induced transcription. Science
308:1789–1791
Smith SE, Read DJ (2008) Mycorrhiza symbiosis, 3rd edn. Academic, San Diego
Soto MJ, Fernandez-Aparicio M, Castellanos-Morales V, Carcia-Garrido JM, Ocampo JA (2010)
First indications for the involvement of strigolactones on nodule formation in alfalfa
(Medicago sativa). Soil Biol Biochem 42:383–385
Valent B, Khang CH (2010) Recent advances in rice blast effector research. Curr Opin Plant Biol
13:434–441
Vandenkoornhuyse P, Leyval C, Bonnin I (2001) High genetic diversity in arbuscular mycorrhizal
fungi: evidence for recombination events. Heredity 87:243–253
Walker SA, Viprey V, Downie JA (2000) Dissection of nodulation signaling using pea mutants
defective for calcium spiking induced in root hairs by Nod factors and chitin oligomers. Proc
Natl Acad Sci USA 97:13413–13418
Weidmann S, Sanchez L, Descombin J, Chatagnier O, Gianinazzi S, Gianinazzi-Pearson V (2004)
Fungal elicitation of signal transduction-related plant genes precedes mycorrhiza establishment and requires the dmi3 gene in Medicago truncatula. Mol Plant Microbe Interact
17:1385–1393
Zwanenburg B, Mwakaboko AS, Reizelman A, Anilkumar G, Sethumadhavan D (2009) Structure
and function of natural and synthetic signalling molecules in parasitic weed germination. Pest
Manag Sci 65:478–491
Signalling and Communication
in the Actinorhizal Symbiosis
Claudine Franche and Didier Bogusz
Abstract More than 200 species of non-legume dicotyledonous plants, mostly
trees and shrubs, belonging to eight different families and 24 genera can enter
actinorhizal symbioses with the nitrogen-fixing actinomycete Frankia. Actinorhizal
nodules consist of multiple lobes, each of which displays a lateral root structure
with infected cells in the expanded cortex. Whereas the key molecules involved in
the molecular dialogue between the symbiotic partners have not yet been
characterized, the development of genomic and molecular tools both in Frankia
and in some actinorhizal plants has contributed to a better understanding of this
original endosymbiosis.
1 Introduction
Actinorhizal plants are a highly diverse group of angiosperms capable of forming
nitrogen-fixing root nodules with the soil actinobacteria Frankia (Huss-Danell
1997; Wall 2000). Phylogenetically, they occur among 25 genera distributed in
eight plant families and four orders with wide-ranging global distributions
(Table 1). Representative species and genera populate temperate forests (Alnus,
Comptonia, Myrica), coastal dunes (Casuarina, Myrica, Hippophae), alpine
biotopes (Alnus, Dryas) and dry areas (Ceanothus, Purshia, Cercocarpus)
(Silvester 1976). Actinorhizal plants share common features; with the exception
of Datisca which has herbaceous shoots, they are perennial dicots and include
woody shrubs and trees. Most actinorhizal plants are capable of high rates of
nitrogen fixation comparable to those found in legumes, and they can sustain
mycorrhizal associations as well (Torrey and Tjepkema 1979; Dawson 2008).
C. Franche (*) • D. Bogusz
Equipe Rhizogénèse, UMR DIADE, Institut de Recherche pour le Développement,
911 avenue Agropolis, BP 64501, 34394 Montpellier Cedex 5, France
e-mail: [email protected]; [email protected]
S. Perotto and F. Baluska (eds.), Signaling and Communication in Plant Symbiosis,
Signaling and Communication in Plants 11, DOI 10.1007/978-3-642-20966-6_4,
# Springer-Verlag Berlin Heidelberg 2012
73
74
C. Franche and D. Bogusz
Table 1 Classification of actinorhizal species and frequency of nodulation within families
(adapted from Dawson 2008)
Plant orders
Family
Nodulated/total genera Genus
Number of species
Fagales
Betulaceae
1/6
Alnus
47
Casuarinaceae 4/4
Allocasuarina
54
Casuarina
16
Ceuthostoma
2
Gymnostoma
18
Myricaceae
2/3
Comptonia
1
Myrica
28
Rosales
Eleagnaceae
3/3
Eleagnus
38
Hippophae
2
Shepherdia
2
Rhamnaceae
8/55
Adolphia
2
Ceanothus
31
Colletia
4
Discaria
5
Kentrothamnus
1
Retanilla
2
Talguenea
1
Trevoa
2
Rosaceae
5/100
Cercocarpus
4
Chamaebatia
1
Cowania
1
Dryas
1
Purshia
2
Cucurbitales Coriariaceae
1/1
Coriaria
5
Datiscaceae
1/3
Datisca
2
Due to these symbiotic properties, actinorhizal plants contribute to improve soil
fertility in disturbed sites and grow well under a range of environmental stresses
such as high salinity, heavy metal pollution and extreme pH. This facility for
adaptation is responsible for the great interest in actinorhizal plants, particularly
in several species of Casuarinaceae such as Casuarina glauca and C. equisetifolia,
which are widely planted in the tropics and subtropics to stabilize sand dunes,
produce fuel wood, rehabilitate area with industrial waste and to increase soil
fertility in agroforestry systems (Dawson 1986; Diem and Dommergues 1990;
Diouf et al. 2008; Zhong et al. 2010).
Our basic knowledge of the symbiotic association between Frankia and
actinorhizal plants is still poorly understood, even though it offers striking
differences with the Rhizobium–legume symbiosis (Pawlowski and Bisseling
1996; Wall 2000; Vessey et al. 2005; Laplaze et al. 2008; Pawlowski and Sprent
2008). Frankia is a filamentous, branching, Gram-positive actinomycete, whereas
Rhizobia are Gram-negative unicellular bacteria. Whereas Rhizobia only enter
symbiosis with plants from the legume family and with one non-legume,
Parasponia, Frankia can interact with a diverse group of non-legume
Signalling and Communication in the Actinorhizal Symbiosis
75
dicotyledonous plants. At the structural level, legume root nodules have peripheral
vascular bundles and infected cells in the central tissue, whereas actinorhizal
nodules display a lateral root structure with a central vascular bundle and peripheral
infected cortical tissue (Duhoux et al. 1996).
The molecular understanding of regulatory events in actinorhizal nodulation is
limited by several factors (Laplaze et al. 2008). One of the major limitations is the
lack of genetic approaches for the host actinorhizal plants that are trees and shrubs.
Additionally, because of their slow growth and high phenolic content, woody plants
such as actinorhizal trees are generally less amenable to physiological and molecular analyses than herbaceous plants. Concerning the microsymbiont Frankia, the
main obstacles are the difficulty to obtain pure culture of the actinobacteria, the
slow-growth rate and the filamentous structure of Frankia which makes it more
difficult to culture in either liquid or solid medium, and the lack of genetic and
mutagenesis tools (Lavire and Cournoyer 2003).
Recently, the development of genomics both in Frankia (Normand et al. 2007a)
and in some actinorhizal plants such as C. glauca (Hocher et al. 2006, 2011),
together with the possibility to obtain transgenic actinorhizal plants following
Agrobacterium gene transfer (Diouf et al. 1995; Franche et al. 1997; Gherbi et al.
2008b; Svistoonoff et al. 2010a), offer new approaches to understand the molecular
basis of the actinorhizal process. In this review, we will highlight recent progress in
the molecular knowledge of the early stages of the actinorhizal symbiosis and
recent data concerning the identification of genes involved in the Frankia factor
(s) signalling pathway. We mainly focus on the interaction between Frankia and the
tropical tree C. glauca, which is one of the most advanced actinorhizal models.
2 The Microorganism Frankia
2.1
General Features of Frankia
The symbiotic microorganism Frankia is the only member of the family
Frankiaceae in the order Actinomycetales (Normand et al. 1996). Frankia is not
an obligate endosymbiont and can occupy two distinct ecological niches, the root
nodule and the soil. Besides its ability to develop a symbiotic interaction with roots,
Frankia is a pleiomorphic bacterium that develops specific structures, vesicules and
spores, that are critical for its survival in the rhizosphere (Newcomb and Wood
1987; Benson and Silvester 1993). Spores are formed in sporangia at the mycelial
tips and are thought to contribute to the dissemination and survival of the microorganism. Vesicules are specialized nitrogen-fixing cells that develop at the tips of
growing vegetative hyphae when exposed to nitrogen starvation and oxygen. They
are surrounded by a laminated lipid envelope containing a mixture of hopanoid
lipids which are pentacyclic bacteriohopanes. The thickness of the vesicle envelope
increases with an increase in O2 and helps to maintain partial oxygen pressure at
76
C. Franche and D. Bogusz
levels that are not labile for the dinitrogenase complex (Parsons et al. 1987; Berry
et al. 1993).
2.2
Specificity of the Symbiotic Association
Since the first report of an infective Frankia strain from nodules of Comptonia
peregrina in 1978 by Callaham et al., hundreds of Frankia isolates have been
described. Individual strains can nodulate actinorhizal plants from different orders,
thus implying that the host origin is not always a determining characteristic for
strain classification. On the basis of cross-infectivity studies and genotypic and
phylogenetic approaches, three cohesive Frankia groups with distinct host ranges
have been defined, as well as a fourth large group of “atypical” strains (Benson and
Clawson 2000; Clawson et al. 2004; Hahn 2008; Normand and Fernandez 2009).
Frankia from Group I include strains which nodulate members of the hamamelid
families Betulaceae, Casuarinaceae and Myricaceae. Frankia from Group II are
typically associated with members of the Coriariaceae, Datiscaceae, Rosaceae and
Ceanothus of the Rhamnaceae. Group II is characterized by low diversity,
supporting the hypothesis of a recent origin for symbiosis in this lineage. Group III
is still poorly known; strains appear to nodulate most Ceanothus sp. and also appear
in Myricaceae, Eleagnaceae, Rosaceae, Betulaceae and Gymnostoma of the
Casuarinaceae family.
Based on their different requirements with respect to Frankia strains,
actinorhizal plants have been divided into promiscuous and non-promiscuous
species (Valdes 2008). Non-promiscuous plant genera include Allocasuarina,
Casuarina, Eleagnus and Hippohae, whereas in greenhouse conditions, Myrica
can be nodulated by most Frankia strains. Although specific, the genera Alnus
and Gymnostoma can sometimes be nodulated by Frankia strains from the
Eleagnaceae infective group. These data suggest that the plant–actinobacterium
interaction could be less specific than the legume–Rhizobium symbiosis.
2.3
Molecular Tools and Genomics of Frankia
The development of genetic tools for Frankia is progressing slowly and remains a
major hurdle for studying the molecular dialogue between the symbiont and the
host plant (Lavire and Cournoyer 2003; Franche et al. 2009). Although there have
been several attempts to develop shuttle vectors from native Frankia plasmids,
there is still no known reliable gene transfer system for the actinomycete (John et al.
2001; Lavire et al. 2001; Xu et al. 2002). Recently, potential transformants of the
strain CcI3 were reported following electroporation with a vector carrying a
tetracyclin resistance gene with a codon usage similar to the one of Frankia, and
driven by a native promoter. Antibiotic selection in liquid culture enabled the
Signalling and Communication in the Actinorhizal Symbiosis
77
growth of tetracyclin-resistant cells (Kucho et al. 2009). However, analysis of the
genomic DNA from the putative transformants revealed that the marker gene was
not integrated in the host. An alternative approach based on the genetic transformation of a uracil-requiring pyrF mutant of CcI3 with a vector carrying the wild-type
pyrF gene is in progress (Kucho et al. unpublished data).
To overcome this limitation and to gain new insights into the nitrogen-fixing
actinobacteria, the genomes of three Frankia strains including CcI3, ACN14a and
EAN1pec, which respectively nodulate C. glauca, Alnus glutinosa and Eleagnus
angustifolia, were compared (Normand et al. 2007a, b; Rawnsley and Tisa 2007).
These genomes were found to be circular and to exhibit high G + C content. The
most striking feature among the three genomes was their sizes, which ranged from
5.43 Mbp for the narrow host range strain CcI3 symbiotic of Casuarina, to 9.04 for
the broad host range strain of Eleagnus EAN1pec. Besides, no significant symbiotic
island or homologues of the common nodABC genes from Rhizobium, which are
required for the synthesis of the core structure of the Nod factors, were identified in
the genomes.
The availability of Frankia genomes has paved the way for the use of bioinformatic approaches such as proteomics. A proteome approach was used on F. alni to
study the physiology of Frankia and to analyse the biochemical events associated
with the switch from N2-replete to N2-fixing conditions (Alloisio et al. 2007). Upregulation of stress proteins and proteins involved in N assimilation was observed
under nitrogen starvation. Additional studies were performed on strain CcI3 to
compare proteome from free-living and symbiotic cells (Mastronunzio et al. 2009).
Solute-binding proteins were the most common class of secreted proteins observed
in the symbiotic proteome, whereas hydrolytic enzymes were rarely detected. These
data suggest that plant cell wall and/or membrane digestion is not likely to be used
by the actinobacteria during the colonization process.
Results of a transcriptome analysis were recently published in the F. alni
ACN14a strain following the expression profiling of 6,607 genes in free-living
nitrogen conditions and symbiotic state (Alloisio et al. 2010). This enabled identification of a large number of up-regulated symbiotic genes such as genes related to
nitrogen fixation, transcriptional regulation, signalling processes, protein drug
export, protein secretion, liposaccharide, and peptidoglycan biosynthesis. Since
strain ACN14a can establish a symbiotic relationship with plants of different
actinorhizal families, the impact of the host plant on the transcriptome of the
symbiont was then studied. Microarray analyses of the transcriptomes in ACN14a
from A. glutinosa, Alnus nepalensis, Myrica gale and Morella rubra did not reveal
any significant differences.
Six more genomes of Frankia are currently being sequenced, and it is expected
that the comparative analysis of these additional genomes with different features
and host specificities will enable a deeper understanding of Frankia in free-living
conditions as well as during different stages of the interaction with hosts of
phylogenetically distant plant families.
78
C. Franche and D. Bogusz
3 The Actinorhizal Nodule
3.1
The Infection Process
Frankia can penetrate the plant root either intracellularly or intercellularly,
depending on the host plant (Fig. 1). In “higher” Hamamelidae such as Alnus,
Myrica, Comptonia, Casuarina and Gymnostoma, the infection process proceeds
through root hair deformation and penetration. The process starts with root hair
curling following the exchange of unknown signals between Frankia and the host
plant (Prin and Rougier 1987). After invagination of growing filaments of Frankia
into the curled root hairs, infection proceeds intracellularly in the root cortex
(Callaham et al. 1979; Berry et al. 1986). Frankia hyphae are surrounded by a
plant-derived encapsulation layer enriched in polygalacturonans, in addition to host
plasmalemma and cytoplasm. This interfacial matrix is the equivalent of the
infection thread wall in legume nodules (Berg 1990, 1999). Upon infection, cell
divisions occur in the root cortex near the infected hairs, leading to a small external
protuberance called a prenodule, which consists of Frankia-infected and uninfected
Fig. 1 Stages of root infection by Frankia. (a) Intracellular infection; (b) Intercellular infection
Signalling and Communication in the Actinorhizal Symbiosis
79
cells (Fig. 2a, b) (Berry and Sunnel 1990). Since the formation of the nodule
primordium does not involve prenodule cells, it is assumed that the prenodule
could be a primitive symbiotic organ (Laplaze et al 2000a). The nodule primordium
results from mitotic activity in pericycle cells located opposite a protoxylem pole
close to the prenodule (Callaham and Torrey 1977; Duhoux et al. 1996). The
infection of the developing nodule appears to be coordinated with the expansion
of post-meristematic cells.
An intercolonization process is observed in 17 genera distributed throughout the
families Eleagnaceae, Rhamnaceae, Rosaceae, Datiscaceae and Coriariaceae,
thus representing 70% of the total actinorhizal species. In this infection pathway,
root hairs are neither deformed nor invaded by the actinobacteria. Frankia enters
the root by growing through the intercellular spaces between adjacent epidermal
cells, penetrating the middle lamella, and progresses apoplastically through cortical
cells within an electron-dense matrix secreted into the intercellular spaces (Miller
and Baker 1985; Liu and Berry 1991; Valverde and Wall 1999; Wall and Berry
2008). Once the nodule primordium has developed from the pericycle, intracellular
penetration by Frankia and the formation of infection threads is initiated acropetally in developing cortical cells of the nodule lobe primordium. In this infection
process, the prenodule stage is not observed. Although the infection process has
been studied in relatively few plant species, it is suggested that the ancestral mode
of infection is via intercellular penetration (Swensen and Benson 2008).
Fig. 2 Actinorhizal nodulation in Casuarina. (a) Prenodule (P) resulting from the infection of
C. glauca by Frankia CcI3. Bar ¼ 1 mm. (b) Longitudinal section of a prenodule showing the
large cortical cells infected by Frankia (F) and the emerging nodular primordium (Pri). Bar ¼ 100
mm. (c) Three-month-old nodules with multiple nodular lobes (Nl) on the root system of
Allocasuarina verticillata. Bar ¼ 1 mm. (d) Three-month-old nodule with nodular lobes (Nl)
and nodular roots (Nr) on the root system of C. glauca. (e) Longitudinal section of an actinorhizal
nodule lobe showing the meristematic zone (Z1), the infection zone (Z2) and the fixation zone
(Z3). Bar ¼ 100 mm. F: Enlargement and lignification of nodule cortical cells resulting from the
infection by Frankia in C. glauca. Bar ¼ 100 mm
80
3.2
C. Franche and D. Bogusz
Nodule Development
Actinorhizal nodules, referred to as actinorhiza, consist of multiple lobes, each of
which resembles a modified lateral root without a cap (Fig. 2c). Nodule lobes have a
central vascular bundle, a superficial periderm and both infected and non-infected
cortical cells. The non-infected cells are thought to play an important role in the
metabolism and transport of the primary product of N2 fixation and assimilation. In
the genera Casuarina, Datisca, Myrica, Comptonia and Gymnostoma, a so-called
root nodule is also observed at the apex of each lobe (Fig. 2d) (Duhoux et al. 1996).
This root nodule lacks root hairs, has a reduced root cap and displays negative
geotropism. It might be involved in the diffusion of gas, especially oxygen, within
and outside the nodule lobe.
Due to activity of the apical meristem, actinorhizal nodule lobes show an
indeterminate growth pattern and have a developmental zonation in which specific
patterns of gene expression are observed (Fig. 2e) (Duhoux et al. 1996). In zone 1,
the apical meristem is free of Frankia. Adjacent to the meristem is an infection zone
where some of the young cortical cells resulting from the meristem activity are
infected by Frankia. The subsequent fixation zone contains both infected and
uninfected cortical cells. Hypertrophied-infected cells are filled with Frankia
hyphae that differentiate N2 fixing vesicles. In some species like Casuarina,
infected cells have a lignified cell wall and no vesicles are observed (Fig. 2f)
(Berg and McDowell 1987). Finally, a basal senescence zone is observed in old
nodules; plant cells and bacteria degenerate and nitrogen fixation is switched off.
Although all nodule lobes have an apical meristem, the growth of individual lobes is
limited. Additional branch lobes are formed as lateral primordia in the pericycle of
the preceding lobe. In plantations of Casuarina, nodules of an average size of 25 cm
have been observed.
4 Early Signal Molecules
4.1
Plant Signals
Actinorhizal nodule development only occurs under conditions of nitrogen deprivation and it is assumed that plant roots emit signals of still unknown nature that are
perceived by Frankia. Root phenylpropanoids are prime candidates for these
signalling molecules, since they govern plant-microorganism cross-talk in the
rhizosphere and regulate the expression of nod genes in Rhizobium (Bhattacharya
et al. 2010). In actinorhizal symbioses, the nodulation of Alnus sp. has been
reported to be enhanced by the addition of flavonone (Benoit and Berry 1997) or
flavonol (Hughes et al. 1999) compounds. There is also some evidence of chemoattraction and proliferation of Frankia in the rhizosphere of Betula pendula, but the
chemical structure of the exuded substances remains unknown (Smolander and
Signalling and Communication in the Actinorhizal Symbiosis
81
Sarsa 1990). In the Myricaceae species, Myrica gale, seed phenolic extracts were
shown to induce general reorganization of protein biosynthesis in several Frankia
strains. Major trends include the up-regulation of oxidative stress proteins such as
FeSOD, which could be involved in protection against host–plant defence
(Bagnarol et al. 2007). In addition, the use of a transcriptomic approach to study
the early stages of the infection process of C. glauca by Frankia revealed that
several genes linked to the flavonoid biosynthesis pathway are up-regulated (Auguy
et al. 2011). The role of flavonoids in the symbiotic process is currently under study
following the down-regulation of the chalcone synthase gene using an RNA
interference approach (Gherbi et al. 2008b) in C. glauca.
4.2
Frankia Signals
In both legumes and actinorhizal plants, the first observable event of the plantmicroorganism dialogue is the curling of the root hairs (Callaham et al. 1979; Van
Ghelue et al. 1997). Preliminary characterization of Frankia deforming factor(s)
suggests that they are structurally divergent from the Nod factors of Rhizobia
(Cérémonie et al 1999). Whereas in legumes Nod factors are heat-stable, amphiphilic
and chitinase sensitive (Dénarié et al. 1996), Frankia factors from the symbiotic
strain ACoN24d of Alnus were found to be heat-stable, hydrophilic, and resistant to
endochitinase and exochitinase from Streptomyces griseus. Moreover, neither the
broad host range Rhizobium sp. NGR234 nor its purified factors elicited root hair
deformation in Alnus and Casuarina, thus suggesting that actinorhizal plants do not
recognize lipo-chito-oligosaccharides. However, N-acetyl-glucosamine, the subunit
of the Nod factor backbone, has been detected in the Alnus root hair deforming
fraction, leaving open the possible existence of a Nod-factor-related Frankia compound (Cérémonie et al. 1999). The lack of a sensitive and reproducible bioassay to
purify Frankia actinorhizal factor(s) remains one of the major obstacles for purifying
the signalling molecules (McEwan et al. 1992).
4.3
Mitotic Activity in Infected Roots
Nod factors have been shown to induce mitotic activity in legume roots (Savouré
et al. 1997). Using Frankia supernatant, no stimulation of mitosis on A. glutinosa
roots was detected (Cérémonie et al. 1999). However, interesting data were obtained
when the cell cycle promoter Pcdc2aAt from Arabidopsis thaliana (Hemerly et al.
1993) was transferred to Casuarina. This gene encodes the P34cdc2 kinase, which is a
key component in the regulation of the cell cycle, acting in the regulation of the G1 to
S and G2 to M transitions (Hemerly et al. 1992; Wang et al. 2004). In non-inoculated
roots of the Casuarinaceae tree Allocasuarina verticillata genetically transformed
with the Pcdc2aAt-GUS construct, the reporter gene expression was mainly observed
82
C. Franche and D. Bogusz
in primary and secondary meristems (Sy et al. 2007). Histological analysis of
b-glucuronidase activity in cdc2aAt-GUS roots, inoculated for 7 days with Frankia,
revealed that cells from the pericycle located opposite the protoxylem poles were
deeply stained. These data suggest that upon Frankia infection, cells from the lateral
roots, and notably the pericycle cells that give rise to nodule primordium, prepare to
re-enter the cell cycle (Sy et al. 2007). It was also observed that adding Frankia
supernatant to cdc2aAt-GUS roots for 2 days contributed to the induction of the GUS
gene, indicating that molecules emitted by Frankia probably stimulate the cell cycle
during the early stages of the interaction, like Nod factors in legumes. However, the
molecules involved in this plant response remain to be characterized.
5 Signal Transduction Pathway
5.1
Functional Characterization of the Candidate Gene SymRK
The development of genetic and genomic tools for the model legumes
M. truncatula and Lotus japonicus has greatly facilitated the cloning of genes
required for root symbiosis (Geurts et al. 2005; Stacey et al. 2006; Oldroyd and
Downie 2008; Madsen et al. 2010). Some of these genes were found to be involved
in the establishment of both rhizobia and mycorrhiza symbioses, and designated as
common Sym genes constituting the common Sym pathway. They include genes
encoding a leucine-rich-repeat (LRR) receptor kinase (SymRK), cation channels,
nuclear pore complex proteins, a calcium and calmodulin-dependent protein kinase
(CCaMK) and a nuclear-coiled protein. The question was raised whether some of
these symbiotic genes were shared in the signal transduction pathway in response to
Frankia factors and rhizobial Nod factors.
To answer this question, a functional study of CgSymRK, a gene isolated from
C. glauca, orthologous to the receptor-like kinase gene SymRK required for nodulation and mycorrhization in legumes, was undertaken (Endre et al. 2002; Stracke
et al. 2002; Capoen et al. 2005). In legumes, this gene (also referred to as DMI2 in
M. truncatula) encodes a leucine-rich-receptor kinase that is needed for bacterial
and endomycorrhizal infection, and Nod-factor-induced calcium spiking in the root
epidermis. Down-regulation of CgSymRK resulting from an RNA interference
approach revealed that the frequency of nodulated RNAi-CgSymRK plants was
reduced twofold compared to control C. glauca plants (Gherbi et al. 2008a). In
addition, a range of morphological alterations was observed in the down-regulated
CgSymRK nodules. Whereas mature nodules in the control plants were multilobed,
RNAi nodules were dramatically reduced in size and mostly consisted in a single
thin lobe that did not fix nitrogen. Cytological analysis further showed that these
aberrant nodular structures accumulated high levels of phenolic compounds and
contained cortical-infected cells that were smaller than untransformed nodular
lobes in controls. Additional experiments revealed that CgSymRK was also
Signalling and Communication in the Actinorhizal Symbiosis
83
necessary for the establishment of the symbiosis with the arbuscular mycorrhiza
Glomus intraradices. The knockdown of CgSymRK was seen to strongly affect
penetration of the fungal hyphae into the root cortex, thus revealing the key role of
CgSymRK in root endosymbioses in Casuarina, and the conservation of SymRK
function between legumes and actinorhizal plants (Gherbi et al. 2008a).
5.2
Search for Other Candidates of the Signalling Pathway
Our group has developed the first genomic platform to identify plant genes involved
in the symbiotic process between Frankia and C. glauca (Hocher et al. 2011). A
total of 2,028 ESTs was first reported from cDNA libraries corresponding to mRNA
extracted from young nodules induced by Frankia and from non-infected roots.
More recently, a new collection of around 40,000 EST corresponding to 15,000
unigenes was obtained from roots and nodulated roots of C. glauca at different
stages of the symbiotic process (Hocher et al. 2011). In half of the nodule
transcripts, no similarity to previously identified genes was detected, while in the
other half, many genes of primary metabolism, protein synthesis, cell division and
defence were identified. To explore the early events of C. glauca-Frankia symbiosis, a substractive hybridization library (SSH) was also constructed with roots
sampled 4 days after infection. A total of 703 SSH sequences were validated and
annotated revealing a large proportion of ESTs involved in defence, cell wall
structure, and gene expression.
Based on the comparison with legume sequences, ESTs sharing significant
homologies with genes from the Nod signalling pathway were identified (Table 2)
(Hocher et al. 2011). Functional analyses of the candidate orthologues to the DMI3
and NIN genes are in progress. DMI3 is a calcium and calmodulin-dependent protein
kinase that is presumed to decode and transduce Nod-factor-specific calcium spiking
Table 2 Nod-factor signalling genes identified in Lotus japonicus (reviewed in Madsen et al.
2010) and homologues EST in Casuarina glauca
L. japonicus
C. glauca ESTs
LysM-RLKs
NFR1
+
NFR5
+
LRR-RLK
SymRK
CgSymRK
Cation channels
POLLUX, CASTOR
+
Nucleoporins
NUP85
NUP133
+
CCaMK
CCaMK
+
GRAS-type TFs
NSP1
+
NSP2
+
TF
NIN
+
The functional characterization of CgSymRK has shown that this actinorhizal gene is functionally
equivalent to SymRK from legumes (Gherbi et al. 2008). +: EST from C. glauca exhibiting
significant homologies with genes from the Nod signalling pathway in legumes. RLK receptorlike kinase, TF transcription factor
84
C. Franche and D. Bogusz
response in legumes (Levy et al. 2004; Mitra et al. 2004). NIN (for nodule inception)
is a transcriptional regulator that is required for the formation of infection threads and
the initiation of nodule primordia. In contrast to DMI3, which is necessary for both
nodulation and endomycorrhization, NIN specifically regulates the development of
root nodules (Schauser et al. 1999; Marsh et al. 2007). From these data, we consequently expect to determine if the symbiotic signalling pathway is conserved in
actinorhizal and rhizobial nitrogen-fixing symbioses beyond the common SYM
genes.
6 Gene Expression During Infection
6.1
Actinorhizal Gene Expression
As expected, the transcriptomic approach has shown that many genes are specifically expressed or enhanced in actinorhizal nodules compared with expression in
roots (Hocher et al. 2006, 2011). Several authors have already reviewed the plant
genes that have been characterized in actinorhizal nodules (Vessey et al. 2005;
Laplaze et al. 2008). Here, we focus on recent data concerning plant genes
expressed in the root hairs and in the infection zone (zone 2) of the nodule.
During the intracellular infection process, the root hair is the primary site of
recognition and infection by the microorganism Frankia (Bhuvaneswari and
Solheim 2000). Since at present no root-hair-enriched cDNA library is available
for any actinorhizal plant, knowledge comes from the characterization of candidate
symbiotic genes using promoter-reporter gene approaches. Plant genes whose
expression is correlated with root hair invasion by the endosymbiont include two
genes from the common Sym pathway, CgSYMRK and CgCCaMK (our group,
unpublished data), and Cg12, a gene from C. glauca encoding a subtilisin-like
serine protease (Laplaze et al. 2000b; Svistoonoff et al. 2003). Subtilases are a
superfamily of proteases that are thought to play a role in different aspects of plant
development including response to pathogens and lateral root development. PCg12
was found to drive expression in infected root hairs and nodule cortical cells
containing growing infection threads. This pattern suggests that the subtilase
CG12 may play a role in the maturation of a polypetide in the infection threads
or in cell wall remodelling resulting from infection by Frankia. Interestingly, the
study of transgenic M. truncatula plants containing the construct PCg12-GUS
construct revealed the same pattern of GUS expression during the infection process
of M. truncatula by Sinorhizobium meliloti (Svistoonoff et al. 2004). It should also
be noted that in absence of a GFP-expressing Frankia strain, transgenic PCg12GUS plants are valuable tools to visualize and count the infection sites during the
early stages of the symbiotic process. The use of this approach revealed that in
PCg12-GUS plants retransformed with an RNAi-CgSymRK construct, the number of
infection sites was considerably reduced, thereby establishing the involvement of
CgSYMRK in root hair infection.
Signalling and Communication in the Actinorhizal Symbiosis
6.2
85
Expression of Heterologous Promoters from Legumes
To investigate the similarities between nitrogen-fixing symbioses in
Rhizobium–legumes and Frankia-actinorhizal plants, heterologous promoters
from early symbiotic genes of legumes were introduced into actinorhizal plants.
Studies concerning the expression pattern conferred by the PGmEnod40-2-GUS
construct showed that the promoter region from the early symbiotic Enod40-2 gene
from soybean was inactive during actinorhizal nodule development (Santi et al.
2003). In legumes, Enod40 genes are first expressed in pericycle cells, and later in
nodule primordium. They play role in nodule initiation and bacteroid development
(Wan et al. 2007).
The heterologous promoter Enod11 from M. truncatula was recently genetically
transformed into C. glauca (Svistoonoff et al. 2010b). In M. truncatula, the
MtEnod11 gene is one of the earliest genes expressed in the root epidermis following
contact with S. meliloti (Journet et al. 2001; Charron et al. 2004). It encodes a putative
cell wall repetitive hydroxy-proline-rich protein and it has proven to be a valuable
marker for early infection-related symbiotic events. In legumes reporter gene activity
driven by the PMtEnod11 promoter was observed in the epidermis within the first
hours of infection by S. meliloti and then in infected root hairs and the infection zone
of young nodules. When introduced into C. glauca, there was no response in the root
epidermis after Frankia inoculation during the root perception stage of the unknown
actinobacterial factors. PMtEnod11-GUS was seen to be expressed as soon as the
actinomycete entered a curled root hair, and subsequently in prenodules and the
nodule cortical cells being infected by Frankia. The expression pattern in nodules
was similar to that conferred by the promoter of PPsENOD12b (Sy et al. 2006),
another early nodulin from Pisum sativum also encoding a repetitive proline-rich
protein (Vijn et al. 1995). From these data, it can be concluded that there is some
conservation of gene regulatory pathways between legumes and actinorhizal plants
in cells involved in bacterial infection and accommodation. Conversely, since there
is no heterologous expression prior to the penetration of Frankia in root hairs, the
perception stages appear to involve transcriptional regulation processes that differ in
actinorhizal plants (Svistoonoff et al. 2010b).
7 Hormones and Actinorhizal Nodulation
7.1
Hormones Produced by Frankia
Bacteria living in association with plants have been shown to produce different
phytohormones that could play a role in the symbiotic interaction. Natural auxins
such as indole-3-acetic acid (IAA) or phenylacetic acid (PAA) were found in
Frankia cultures at relatively high concentrations (105 to 106 M) (Wheeler and
Henson 1979; Wheeler et al. 1984; Hammad et al. 2003). The cytokinin isopentenyl
86
C. Franche and D. Bogusz
adenosine (iPA) was also detected at a concentration of 106 M (Gordons et al.
1988; Stevens and Berry 1988). However, it remains to be established whether or
not Frankia produces auxin and/or cytokinin in planta during the infection process.
Transcriptome analyses of the symbiotic form of the actinobacterial genes will
likely provide some evidence to elucidate the contribution of the symbiont in the
hormonal content of the nodule.
7.2
A Role for Auxin
Auxins are a class of plant growth hormones with many roles in cell division and
enlargement, differentiation, lateral root formation, and vascular bundle formation
(Casimiro et al. 2003). Since these biological processes occur during actinorhizal
nodule formation and because actinorhizal nodules anatomically and ontogenetically resemble lateral roots, the role of auxin in the actinorhizal symbiosis was
addressed. Elevated levels of auxins were measured in the actinorhizae (Wheeler
and Henson 1979) and application of auxin efflux transport inhibitors such as NPA
or TIBA on C. glauca roots led to the formation of nodule-like structures called
“pseudonodules” on the roots of Casuarinaceae plantlets (Duhoux et al. 1996).
Attempts to use auxin-sensitive promoters such as the soybean GH3 promoter
(Reddy et al. 2006) or the synthetic DR5 promoter (Ulmasov et al. 1995) to monitor
changes either in auxin concentration or in cellular sensitivity during the infection
process were unsuccessful.
However, interesting results resulted from the functional analysis of the
CgAUX1, which encodes a high-affinity auxin influx transporter in C. glauca.
Using the GUS gene driven by the CgAUX1 promoter, it was clearly shown that
auxin plays an important role during plant cell infection in actinorhizal symbiosis
(Péret et al. 2007). CgAUX1-GUS was found to be strongly expressed both during
root hair infection and cortical cell invasion by Frankia. It was also reported that
CgAUX1-GUS was highly expressed in the root primordium, whereas no reporter
gene activity was observed in the nodule primordium. This result further indicates
that, although the actinorhizal nodule is comparable with a symbiotic lateral root,
the molecular mechanisms involved in primordia initiation in lateral roots may
differ from those in actinorhizal nodules.
7.3
Cytokinin and Actinorhizal Nodulation
Besides auxin, elevated levels of cytokinins have been found in actinorhizal
nodules (Wheeler and Henson 1979) and the induction of bacteria-free nodules,
so-called pseudo-actinorhiza, by cytokinin has been reported (Rodriguez-Barrueco
and de Castro 1973). To monitor changes in cytokinin levels during the actinorhizal
nodulation process, we introduced into Casuarina a transcriptional fusion between
Signalling and Communication in the Actinorhizal Symbiosis
87
the reporter gene GUS and the Arabidopsis cytokinin-responsive gene ARR5
(D’Agostino et al. 2000). b-Glucuronidase activity was found to be restricted to
the root cap and the root meristems, and no change in expression resulted from
infection by Frankia, thus implying that this heterologous promoter was not
appropriate to study changes in cytokinin levels in Casuarina. More valuable data
will probably come from the isolation of cytokinin-responsive genes from
actinorhizal plants.
8 Conclusions
Although several actinorhizal genera contain species that are economically important in forestry and land regeneration (Dawson 1986), progress has been slower in
our knowledge of the actinorhizal symbiosis than that of the legume–Rhizobium
interaction. Major advances have nevertheless been made in recent years thanks to
the development of genomics and the transgenic tools. Progress includes the
sequencing and comparative analysis of an increasing number of Frankia genomes,
even though the most decisive progress will come from the development of a
genetic approach in Frankia. The availability of a method for the genetic transformation of Casuarinas has resulted in a major breakthrough since it opened the way
to functional gene analysis in the actinorhizal host plant. It recently led to a major
result establishing that the unknown Frankia factors, Rhizobial Nod factors and
Myc factors use a common pathway (Gherbi et al. 2008a). Deep sequencing
analyses are now in progress in actinorhizal plants and mining these data will
undoubtly contribute to dissecting the molecular dialogue between Frankia and
the host plant in the different stages of development of the actinorhizal nodule.
References
Alloisio N, Felix S, Marechal J, Pujic P, Rouy Z, Vallenet D, Medigue C, Normand P (2007)
Frankia alni proteome under nitrogen-fixing and nitrogen-replete conditions. Physiol Plant
130:440–453
Alloisio N, Queiroux C, Fournier P, Pujic P, Normand P, Vallenet D, Médigue C, Yamaura M,
Kakoi K, Kucho K-I (2010) The Frankia alni symbiotic transcriptome. Mol Plant Microbe
Interact 23:593–607
Auguy F, Abdel-Lateif K, Doumas P, Badin P, Guerin V, Bogusz D, Houcher V (2011)
Isoflavonoids pathway activation in actinorhizal symbioses. Funct Plant Biol, in press
Bagnarol E, Popovici J, Alloisio N, Maréchal J, Pujic P, Normand P, Fernandez M (2007)
Differential Frankia protein patterns induced by phenolic extracts from Myricaceae seeds.
Physiol Plant 130:380–390
Benoit LF, Berry AM (1997) Flavonoid-like compounds from red alder (Alnus rubra) influence
nodulation by Frankia (Actinomycetales). Physiol Plant 99:588–593
88
C. Franche and D. Bogusz
Benson DR, Clawson ML (2000) Evolution of the actinorhizal plant symbioses. In: Triplett EW
(ed) Prokaryotic nitrogen fixation: a model system for analysis of biological process. Horizon
Scientific Press, Wymondham, UK, pp 207–224
Benson DR, Silvester WB (1993) Biology of Frankia strains, actinomycete symbionts of
actinorhizal plants. Microbiol Rev 57:293–319
Berg RH (1990) Cellulose and xylans in the interface capsule in symbiotic cells of actinorhizae.
Protoplasma 159:35–43
Berg RH (1999) Frankia forms infection threads. Can J Bot 77:1327–1333
Berg RH, McDowell L (1987) Endophyte differentiation in Casuarina actinorhizae. Protoplasma
136:104–117
Berry AM, Sunnel LA (1990) The infection process and nodule development. In: Schwintzer CR,
Tjepkema JD (eds) The biology of Frankia and actinorhizal plants. Academic, New York,
pp 61–81
Berry AM, McIntyre L, McCully M (1986) Fine structure of root hair infection leading to
nodulation in the Frankia-Alnus symbiosis. Can J Bot 64:292–305
Berry AM, Harriott OT, Moreau RA, Osman SF, Benson DR, Jones AD (1993) Hopanoid lipids
compose the Frankia vesicle envelope, presumptive barrier of oxygen diffusion to nitrogenase.
Proc Natl Acad Sci USA 90:6091–6094
Bhattacharya A, Sood P, Citovsky V (2010) The roles of plant phenolics in defense and communication during Agrobacterium and Rhizobium infection. Mol Plant Pathol 11:705–719
Bhuvaneswari TV, Solheim B (2000) Root-hair interactions in actinorhizal symbioses. In: Ridge RW,
Emons AMC (eds) Root hairs – cell and molecular biology. Springer, Heidelberg, pp 311–327
Callaham D, Torrey JG (1977) Prenodule formation and primary nodule development in roots of
Comptonia (Myricaceae). Can J Bot 51:2306–2318
Callaham D, DelTredici P, Torrey JG (1978) Isolation and cultivation in vitro of the actinomycete
causing root nodulation in Comptonia. Science 199:899–902
Callaham D, Newcomb W, Torrey JG, Peterson RL (1979) Root hair infection in actinomyceteinduced root nodule initiation in Casuarina, Myrica and Comptonia. Bot Gaz 140:S1–S9
Capoen W, Goormachtig S, De Rycke R, Schoeyers K, Holsters M (2005) SrSymRK, a plant
receptor essential for symbiosome infection. Proc Natl Acad Sci USA 102:10369–10374
Casimiro I, Beeckman T, Graham N, Bhalerao R, Zhang H, Casero P, Sandberg G, Bennett MJ
(2003) Dissecting Arabidopsis lateral root development. Trends Plant Sci 8:165–171
Cérémonie H, Debellé F, Fernandez MP (1999) Structural and functional comparison of Frankia
root hair deforming factor and rhizobia Nod factor. Can J Bot 77:1293–1301
Charron D, Pingret JL, Chabaud M, Journet EP, Barker DG (2004) Pharmacological evidence that
multiple phospholipid signaling pathways link Rhizobium nodulation factor perception in
Medicago truncatula root hairs to intracellular responses, including Ca2+ spiking and specific
ENOD gene expression. Plant Physiol 136:3582–3593
Clawson ML, Bourret A, Benson DR (2004) Assessing the phylogeny of Frankia-related plant
nitrogen-fixing root nodule symbioses with Frankia 16SRNA and glutamine synthetase gene
sequences. Mol Phylogenet Evol 31:131–138
D’Agostino IB, Deruère J, Kieber JJ (2000) Characterization of the response of the arabidopsis
response regulator gene family to cytokinin. Plant Physiol 124:1706–1717
Dawson JO (1986) Actinorhizal plants: their use in forestry and agriculture. Outlook Agric 15
(4):202–208
Dawson JO (2008) Ecology of actinorhizal plants. In: Pawlowski K, Newton WE (eds) Nitrogen
fixation research: origins and progress, vol VI, Nitrogen-fixing Actinorhizal symbioses.
Springer, Heidelberg, pp 199–234
Dénarié J, Debellé F, Promé JC (1996) Rhizobium lipochitooligosaccharide nodulation factor:
signaling molecules mediating recognition and morphogenesis. Annu Rev Biochem 65:503–535
Diem HG, Dommergues YR (1990) Current and potential uses and management of Casuarinaceae
in the tropics and subtropics. In: Schwintzer R, Tjepkema JD (eds) The biology of Frankia and
actinorhizal plants. Academic, San Diego, pp 317–342
Signalling and Communication in the Actinorhizal Symbiosis
89
Diouf D, Gherbi H, Prin Y, Franche C, Duhoux E, Bogusz D (1995) Hairy root nodulation of
Casuarina glauca: a system for the study of symbiotic gene expression in an actinorhizal tree.
Mol Plant Microbe Interact 8:532–537
Diouf D, Sy M-O, Gherbi H, Bogusz D, Franche C (2008) Casuarinaceae. In: Kole CR, Scorza R,
Hall TC (eds) Compendium of transgenic crop plants, vol 9, Transgenic forest tree species.
Blackwell, Oxford, UK, pp 279–292
Duhoux E, Diouf D, Gherbi H, Franche C, Ahée J, Bogusz D (1996) Le nodule actinorhizien. Acta
bot Gallica 143:593–608
Endre G, Kereszt A, Devei Z, Mihacea S, Kalo P, Kiss G (2002) A receptor kinase gene regulating
symbiotic nodule development. Nature 417:962–966
Franche C, Diouf D, Le QV, N’Diaye A, Gherbi H, Bogusz D, Gobé C, Duhoux E (1997) Genetic
transformation of the actinorhizal tree Allocasuarina verticillata by Agrobacterium
tumefaciens. Plant J 11:897–904
Franche C, Lindstr€om K, Elmerich C (2009) Nitrogen-fixing bacteria associated with leguminous
and non-leguminous plants. Plant Soil 321:35–59
Geurts R, Federova E, Bisseling T (2005) Nod factor signaling genes and their function in the early
stages of Rhizobium infection. Curr Opin Plant Biol 8:346–352
Gherbi H, Markmann K, Svistoonoff S, Estevan J, Autran D, Giczey G, Auguy F, Péret B, Laplaze L,
Franche C, Parniske M, Bogusz D (2008a) SymRK defines a common genetic basis for plant root
endosymbioses with AM fungi, rhizobia and Frankia bacteria. Proc Natl Acad Sci USA
105:4928–4932
Gherbi H, Nambiar-Veetil M, Zhong C, Félix J, Autran D, Girardin R, Vaissayre V, Auguy F,
Bogusz D, Franche C (2008b) Post-transcriptional gene silencing in the root system of the
actinorhizal tree Allocasuarina verticillata. Mol Plant Microbe Interact 21:518–524
Gordons A, Stevens JR, Berry AM (1988) Cytokinin secretion by Frankia sp. HFPArI3 in defined
medium. Plant Physiol 87:15–16
Hammad Y, Nalin R, Marechal J, Fiasson K, Pepin RAM, Normand P, Domenach AM (2003) A
possible role for phenyl acetic acid (PAA) on Alnus glutinosa nodulation by Frankia. Plant Soil
254:193–205
Hann D (2008) Polyphasic taxonomy of the genus Frankia. In: Pawlowski K, Newton WE (eds)
Nitrogen fixation research: origins and progress, vol VI, Nitrogen-fixing actinorhizal
symbioses. Springer, Heidelberg, pp 25–47
Hemerly AS, Bergouniou C, Van Montagu M, Inzé D, Ferreira P (1992) Gene regulating the plant
cell cycle: isolation of a mitotic-like cyclin from Arabidodpsis thaliana. Proc Natl Acad Sci
USA 89:3295–3299
Hemerly AS, Ferreira PCG, de Almeida EJ, van Montagu M (1993) cdc2a expression in
Arabidopsis thaliana is linked with competence for cell division. Plant Cell 5:1711–1723
Hocher V, Auguy F, Argout X, Laplaze L, Franche C, Bogusz D (2006) Expressed sequence – tag
analysis in Casuarina glauca actinorhizal nodule and root. New Phytol 169:681–688
Houcher V, Alloisio N, Auguy F, Fournier P, Doumas P, Pujic P, Gherbi H, Queiroux C,
Da Silva C, Wincker P, Normand P, Bogusz D (2011) Transcriptomics of actionorhizal
symbioses reveals homologs of the whole common symbiotic signaling cascade. Plant Physiol
56:700–771
Hughes M, Donnelly C, Crozier A, Wheeler CT (1999) Effects of the exposure of roots of Alnus
glutinosa to light on flavonoids and nodulation. Can J Bot 77:1311–1315
Huss-Danell K (1997) Actinorhizal symbioses and their N2 fixation. New Phytol 136:375–405
John TR, Rice JM, Johnson JD (2001) Analysis of pFQ12, a 22.4-kb Frankia plasmid. Can J
Microbiol 47:608–617
Journet EP, El-Gachtouli N, Vernoud V, de Billy F, Pichon M, Dedieu A, Arnould C, Morandi D,
Barker DG, Gianinazzi-Pearson V (2001) Medicago truncatula ENOD11: a novel RPRPencoding early nodulin gene expressed during mycorrhization in arbuscule-containing cells.
Mol Plant Microbe Interact 14:737–748
90
C. Franche and D. Bogusz
Kucho K-I, Kakoi K, Yamaura M, Higashi S, Uchiumi T, Abe M (2009) Transient transformation
of Frankia by fusion marker genes in liquid culture. Microbes Environ 24:231–240
Laplaze L, Duhoux E, Franche C, Frutz T, Svistoonoff S, Bisseling T, Bogusz D, Pawlowski K
(2000a) Casuarina glauca prenodule cells display the same differentiation as the
corresponding nodule cells. Mol Plant Microbe Interact 13:107–112
Laplaze L, Ribeiro A, Franche C, Duhoux E, Auguy F, Bogusz D, Pawlowski K (2000b)
Characterization of a Casuarina glauca nodule-specific subtilisin-like protease gene, a homolog of Alnus glutinosa ag12. Mol Plant Microbe Interact 13:113–117
Laplaze L, Svistoonoff S, Santi C, Auguy F, Franche C, Bogusz D (2008) Molecular biology of
actinorhizal symbioses. In: Pawlowski K, Newton WE (eds) Nitrogen fixation research: origins
and progress, vol VI, Nitrogen-fixing actinorhizal symbioses. Springer, Heidelberg, pp
235–259
Lavire C, Cournoyer B (2003) Progress on the genetics of the N2-fixing actinorhizal symbiont
Frankia. Plant Soil 254:125–137
Lavire C, Louis D, Perriere G, Briolay J, Normand P, Cournoyer B (2001) Analysis of pFQ31, a
8551-bp cryptic plasmid from three symbiotic nitrogen-fixing actinomycete Frankia. FEMS
Microbiol Lett 197:111–116
Levy J, Bres C, Geurts R, Chalhoub B, Kulikova O, Duc G, Journet E, AnéJ-M LE, Bisseling T,
Dénarié J, Rosenberg C, Debellé F (2004) A putative Ca2+ and calmodulin-dependent protein
kinase required for bacterial and fungal symbioses. Science 303:1361–1364
Liu Q, Berry AM (1991) The infection process and nodule initiation in the Frankia-Ceanothus root
nodule symbiosis. Protoplasma 163:82–92
Madsen LH, Tirichine L, Jurkiewicz A, Heckmann AB, Bek AS, Ronson CW, James EK,
Stougaard J (2010) The molecular network governing nodule organogenesis and infection in
the model legume Lotus japonicus. Nat commun 1:1–12
Marsh JF, Rakocevic A, Mitra RM, Brocard L, Sun J, Eschstruth A, Logn SR, Schultze M, Ratet P,
Oldroyd GED (2007) Medicago truncatula NIN is essential for rhizobial-independent nodule
organogenesis induced by autoactive calcium/calmodulin-dependent protein kinase. Plant
Physiol 144:324–335
Mastronunzio JE, Huang Y, Besnon DR (2009) Diminished exoproteome of Frankia spp. in
culture and symbiosis. App Environ Microbiol 75:6721–6728
McEwan NR, Green DC, Wheeler CT (1992) Utilisation of the root hair curling reaction in Alnus
glutinosa for the assay of nodulation signal compounds. Acta Oecol 13:509–510
Miller IM, Baker DD (1985) The initiation, development and structure of root nodules in Elaegnus
angustifolia L. (Elaeagnaceae). Protoplasma 128:107–119
Mitra RM, Gleason CA, Edwards A, Hadfield J, Downie JA, Olroyd GE, Long SR (2004) A Ca2+/
calmodulin dependent protein kinase required for symbiotic nodule development: gene identification by transcript-based cloning. Proc Natl Acad Sci USA 101:4701–4705
Newcomb WR, Wood S (1987) Morphogenesis and fine structure of Frankia (Actinomycetales):
the microsymbiont of nitrogen-fixing actinorhizal root nodules. Int Rev Cytol 109:1–88
Normand P, Fernandez MP (2009) Evolution and diversity of Frankia. Microbiol Monogr
8:103–125
Normand P, Orso S, Cournoyer B, Jeannin P, Chapelon C, Dawson J, Evtushenko L, Misra AK
(1996) Molecular phylogeny of the genus Frankia and related genera and emendation of family
Frankiaceae. Int J Syst Bacteriol 46:1–9
Normand P, Lapierre P, Tisa LS, Gogarten JP, Alloisio N, Bagnarol E, Bassi CA, Berry AM,
Bickhart DM, Choisne N, Couloux A, Cournoyer B, Cruveiller S, Daubin V, Demange N,
Francino MP, Goltsman E, Huang Y, Martinez M, Mastronunzio JE, Mullin BC, Nieman J,
Pujic P, Rawnsley T, Rouy Z, Schenowitz C, Sellstedt A, Tvares F, Tomkins JP, Vallenet D,
Valverde C, Wall L, Wang Y, Medigue C, Benson DR (2007a) Genome characteristics of
facultatively symbiotic Frankia sp. strains reflect host range and host plant biogeography.
Genome Res 17:7–15
Signalling and Communication in the Actinorhizal Symbiosis
91
Normand P, Queiroux C, Tisa LS, Benson DR, Rouy Z, Cruveiller S, Medigue C (2007b)
Exploring the genomes of Frankia. Physiol Plant 130:331–343
Oldroyd GED, Downie JA (2008) Coordinating nodule morphologenesis with rhizobial infection
in legumes. Annu Rev Plant Biol 59:519–546
Parsons R, Silvester WB, Harris S, Gruijters WTM, Bullivant S (1987) Frankia vesicles provide
inducible and absolute oxygen protection for nitrogenase. Plant Physiol 83:728–731
Pawlowski K, Bisseling T (1996) Rhizobial and actinorhizal symbioses: what are the shared
features? Plant Cell 8:1899–1913
Pawlowski K, Sprent J (2008) Comparison between actinorhizal and legume symbiosis. In:
Pawlowski K, Newton WE (eds) Nitrogen-fixing actinorhizal symbioses. Springer, Dordrecht,
The Netherlands, pp 261–288
Péret B, Swarup R, Jansen L, Devos G, Auguy F, Collin M, Santi C, Hocher V, Franche C, Bogusz
D, Bennett M, Laplaze L (2007) Auxin influx activity is associated with Frankia infection
during actinorhizal nodule formation in Casuarina glauca. Plant Physiol 144:1852–1862
Prin Y, Rougier M (1987) Preinfection events in the establishment of Alnus-Frankia symbiosis:
study of the root hair deformation step. Plant Physiol 6:99–106
Rawnsley T, Tisa LS (2007) Development of a physical map for three Frankia strains and a partial
genetic map for Frankia EuI1c. Physiol Plant 130:427–439
Reddy SM, Hitchin S, Melayah D, Pandey AK, Raffier C, Henderson J, Marmeisse R, Gay G
(2006) The auxin-inducible GH3 homologue Pp-GH3.16 is downregulated in Pinus pinaster
root systems on ectomycorrhizal symbiosis establishment. New Phytol 170:391–400
Rodriguez-Barrueco F, De Castro B (1973) Cytokinin-induced pseudonodules in Alnus glutinosa.
Physiol Plant 29:277–280
Santi C, von Groll U, Ribeiro A, Chiurazzi M, Auguy F, Bogusz D, Franche C, Pawlowski K
(2003) Comparison of nodule induction in legume and actinorhizal symbiosis: the induction of
actinorhizal nodules does not involve ENOD40. Mol Plant Microbe Interact 16:808–816
Savouré A, Sallaud C, El-Turk J, Zuanazzi J, Ratet P, Schultze M, Kondorosi A, Esnault R,
Kondorosi E (1997) Distinct response of Medicago suspension cultures ad roots to Nod factors
and chitin oligomers in the elicitation of defense-related responses. Plant J 11:277–287
Schauser L, Roussis A, Stiller J, Stougaard J (1999) A plant regulator controlling development of
symbiotic root nodules. Nature 402:191–195
Silvester WB (1976) Ecological and economic significance of the non-legume symbiosis. In:
Newton WE, Nyman CJ (eds) Proceedings first international symposium on nitrogen fixation.
Washington University Press, Pullman
Smolander A, Sarsa M-L (1990) Strains of soil under Betula pendula: behaviour in soil and pure
culture. Plant Soil 122:129–136
Stacey G, Libault M, Brechenmacher L, Wan J, May G (2006) Genetics and functional genomics
of legume nodulation. Curr Opin Plant Biol 9:110–121
Stevens GA, Berry AM (1988) Cytokinin secretion by Frankia sp. HFPAr13 in defined medium.
Plant Physiol 87:15–16
Stracke S, Kistner C, Yoshida S, Mulder L, Sato S, Kaneko T, Tabata S, Sandal N, Stougaard J,
Szczyglowski K (2002) A plant receptor-like kinase required for both bacterial and fungal
symbiosis. Nature 417:959–962
Svistoonoff S, Laplaze L, Auguy F, Runions CJ, Duponnois R, Haseloff J, Franche C, Bogusz D
(2003) Cg12 expression is specifically linked to infection of root hairs and cortical cells during
Casuarina glauca and Allocasuarina verticillata actinorhizal nodule development. Mol Plant
Microbe Interact 16:600–607
Svistoonoff S, Laplaze L, Liang J, Ribeiro A, Gouveia MC, Auguy F, Fevereiro P, Franche C,
Bogusz D (2004) Infection-related activation of the Cg12 promoter is conserved between
actinorhizal and legume-rhizobia root nodule symbiosis. Plant Physiol 136:3191–3197
Svistoonoff S, Gherbi H, Nambiar-Veetil M, Zhong C, Michalak Z, Laplaze L, Vaissayre V,
Auguy F, Hocher V, Doumas P, Bonneau J, Bogusz D, Franche C (2010a) Contribution of
transgenic Casuarinaceae to our knowledge of the actinorhizal symbioses. Symbiosis 50:3–11
92
C. Franche and D. Bogusz
Svistoonoff S, Sy M-O, Diagne N, Barker D, Bogusz D, Franche C (2010b) Infection-specific
activation of the Medicago truncatula Enod11 early nodulin gene promoter during actinorhizal
root nodulation. Mol Plant Microbe Interact 23:740–747
Swensen SM, Benson DR (2008) Evolution of actinorhizal host plants and Frankia endosymbionts.
In: Pawlowski K, Newton WE (eds) Nitrogen fixation research: origins and progress, vol VI,
Nitrogen-fixing Actinorhizal symbioses. Springer, Heidelberg, pp 73–104
Sy M-O, Constans L, Obertello M, Geney C, Laplaze L, Auguy F, Hocher V, Bogusz D, Franche C
(2006) Analysis of the expression pattern conferred by the PsEnod12B promoter from the early
nodulin gene of Pisum sativum in transgenic actinorhizal trees of the Casuarinaceae family.
Plant Soil 281:281–289
Sy M-O, Hocher V, Gherbi H, Laplaze L, Auguy F, Bogusz D, Franche C (2007) The cell-cycle
promoter cdc2aAt from Arabidopsis thaliana is induced in the lateral roots of the actinorhizal
tree Allocasuarina verticillata during the early stages of the symbiotic interaction with
Frankia. Physiol Plant 130:409–417
Torrey JG, Tjepkema JD (1979) Symbiotic nitrogen fixation in actinomycete-nodulated plants. Bot
Gaz 140(Suppl):i–ii
Ulmasov T, Liu ZB, Hagen G, Guilfoyle TJ (1995) Composite structure of auxin response
elements. Plant Cell 7:1611–1623
Valdes M (2008) Frankia ecology. In: Pawlowski K, Newton WE (eds) Nitrogen fixation research:
origins and progress, vol VI, Nitrogen-fixing actinorhizal symbioses. Springer, Heidelberg, pp
49–72
Valverde C, Wall LG (1999) Time course of nodule development in Discaria trinervis
(Rhamnaceae) – Frankia symbiosis. New Phytol 141:345–354
van Ghelue M, Lovaas E, Ringo E, Solheim B (1997) Early interaction between Alnus glutinosa
and Frankia strain Arl3. Production and specificity of root hair deformation factors. Physiol
Plant 99:579–587
Vessey JK, Pawlowski K, Bergman B (2005) Root-based N2-fixing symbioses: legumes,
actinorhizal plants, Parasponia sp. and cycads. Plant Soil 274:51–78
Vijn I, Christiansen H, Lauridsen P, Kardailsky I, Quandt H-J, Broer II, Drenth J, Ostergaard E,
van Kammen A, Bisseling T (1995) A 200 bp region of the pea ENOD12 promoter is sufficient
for nodule-specific and nod factor induced expression. Plant Mol Biol 28:1103–1110
Wall LG (2000) The actinorhizal symbiosis. J Plant Growth Regul 19:167–182
Wall LG, Berry AM (2008) Early interactions, infection and nodulation in actinorhizal symbiosis.
In: Pawlowski K, Newton WE (eds) Nitrogen-fixing actinorhizal symbioses. Springer,
Dordrecht, The Netherlands, pp 147–166
Wan X, Hontelez J, Lillo A, Guarnerio C, van de Peut D, Fedorova E, Bisseling T, Franssen H
(2007) Medicago truncatula ENOD40-1 and ENOD40-2 are both involved in nodule initiation
and bacteroid development. J Exp Bot 58:2033–2041
Wang G, Kong H, Sun Y, Zhang X, Zhang W, Altman N, De Pamphilis CW, Ma H (2004)
Genomic wide analysis of the cyclin family in Arabidopsis and comparative phylogenetic
analysis of plant cyclin-like proteins. Plant Physiol 135:1084–1099
Wheeler CT, Henson IE (1979) Hormones in plants bearing actinomycete nodules. Bot Gaz
140:52–57
Wheeler CT, Crozier A, Sandberg G (1984) The biosynthesis of indole-3 acetic acid by Frankia.
Plant Soil 78:99–104
Xu XD, Kong RQ, de Bruijn FJ, He SY, Murry MA, Newman T, Wolk P (2002) DNA sequence
and genetic characterization of plasmid pFQ11 from Frankia alni strain CpI1. FEMS
Microbiol Lett 207:103–107
Zhong C, Zhang Y, Chen Y, Jiang Q, Chen Z, Liang J, Pinyopusarek K, Franche C, Bogusz D
(2010) Casuarina research and applications in China. Symbiosis 50:107–114
Signalling in Cyanobacteria–Plant Symbioses
David G. Adams and Paula S. Duggan
Abstract Cyanobacteria are a morphologically diverse and widespread group of
phototrophic bacteria, many of which are capable of nitrogen fixation. They form
symbioses with a wide range of eukaryotic hosts including fungi (lichens and
Geosiphon pyriformis), diatoms, dinoflagellates, sponges, ascidians (sea squirts),
corals and plants. The best understood are the plant symbioses, which are the subject
of this chapter. In the cyanobacteria–plant associations, the cyanobacteria provide the
host with fixed nitrogen and usually adopt a heterotrophic form of nutrition, using
fixed carbon supplied by the plant, enabling them to occupy regions of the host, such
as the roots, that receive little or no light. Most cyanobacterial symbionts of plants
belong to the genus Nostoc, members of which fix nitrogen in specialised cells known
as heterocysts, which provide the necessary microoxic environment for the functioning of the oxygen-sensitive enzyme nitrogenase. These cyanobacteria, which are
immotile for most of their life cycles, produce specialised motile filaments known as
hormogonia, as a means of dispersal and as the infective agents in plant symbioses.
Host plants improve their chances of infection by releasing external chemical signals
that both stimulate hormogonia formation and serve as chemoattractants. However,
within the symbiotic tissue the plant releases hormogonia-repressing factors to ensure
the conversion of hormogonia into heterocyst-containing, nitrogen-fixing filaments.
1 Introduction
Cyanobacteria are a large group of morphologically diverse phototrophic bacteria
found in almost every environment on Earth. Because of their abundance, especially in the oceans, they make enormous contributions to global carbon and
D.G. Adams (*) • P.S. Duggan
Faculty of Biological Sciences, Institute of Integrative and Comparative Biology,
University of Leeds, Leeds LS2 9JT, UK
e-mail: [email protected]; [email protected]
S. Perotto and F. Baluska (eds.), Signaling and Communication in Plant Symbiosis,
Signaling and Communication in Plants 11, DOI 10.1007/978-3-642-20966-6_5,
# Springer-Verlag Berlin Heidelberg 2012
93
94
D.G. Adams and P.S. Duggan
nitrogen fixation. Cyanobacteria form symbiotic associations with a wide variety of
eukaryotic hosts including plants, fungi, sponges and protists. The cyanobacterial
symbionts (cyanobionts) can provide fixed carbon for non-photosynthetic hosts
such as the fungi of both lichens (Rikkinen 2002) and Geosiphon pyriformis
(Kluge 2002; Kluge et al. 2002; Adams et al. 2006), but their primary role is to
provide fixed nitrogen, enabling the host to occupy nitrogen-poor environments.
They may also protect some hosts, such as ascidians and isopod crustaceans, from
grazing by predators, and others, such as sponges, from excessive sunlight (Usher
2008). The benefits to the cyanobionts are less clear but are likely to include
protection from competition, predation and environmental extremes. In the case
of plants, the cyanobionts can also adapt to a heterotrophic life style using carbon
from the plant host, enabling them to devote more resources to the fixation of
nitrogen for both partners, and to grow in plant structures, such as the roots of
cycads and the stem glands of Gunnera that receive little or no light.
Cyanobacteria vary in morphology from simple unicells to complex filamentous
forms capable of differentiating several specialised cell types including the nitrogen-fixing heterocyst (Fig. 7c, d) which provides suitable microoxic conditions for
the fixation of nitrogen by the oxygen-sensitive enzyme nitrogenase (Meeks et al.
2002; Meeks and Elhai 2002; Golden and Yoon 2003; Zhang et al. 2006; Flores and
Herrero 2010). The cyanobionts of plants are filamentous heterocyst-forming
cyanobacteria primarily from the genus Nostoc. Another important characteristic
of these cyanobacteria is the production of specialised, motile filaments known as
hormogonia. These provide Nostoc spp. (and cyanobacteria of other heterocystous
genera such as Calothrix and Fischerella) with a motile phase and hence a means of
dispersal in an otherwise sessile life cycle. Hormogonia are also the infective agent
in the plant symbioses (Meeks et al. 2002; Meeks and Elhai 2002; Meeks 2009;
Flores and Herrero 2010). Indeed, many plants release unidentified chemicals that
stimulate the formation of hormogonia and attract them to the symbiotic tissues
(Sect. 3).
2 The Cyanobacteria–Plant Symbioses
This chapter deals with the endophytic cyanobacteria–plant symbioses in which the
cyanobiont is located within the host tissue or, in the case of the angiosperm
Gunnera, within the host cells. However, many cyanobacteria, including nitrogen-fixing strains, grow as epiphytes on a wide range of plants, particularly in
aquatic environments (Adams 2011). These “loose” associations won’t be
discussed here, as they are poorly studied and little is known about the extent to
which the plant benefits, although the nitrogen-fixing cyanobacteria certainly contribute to the local nitrogen economy. By contrast, in the endophytic associations
the plant host clearly benefits from the provision of combined nitrogen by the
cyanobionts and there are clear, although not well understood, interactions between
the cyanobiont and its host.
Signalling in Cyanobacteria–Plant Symbioses
2.1
95
Bryophytes (Mosses, Hornworts and Liverworts)
The mosses, hornworts and liverworts are small, non-vascular land plants (Figs. 1
and 3a), some of which can become infected with primarily heterocystous
cyanobacteria (Adams 2002a, b; Meeks 2003; Solheim et al. 2004; Adams et al.
2006; Adams and Duggan 2008; Bergman et al. 2007a, 2008). The cyanobacteria in
moss associations are mostly epiphytic (Solheim and Zielke 2002; Solheim et al.
2004; Gentili et al. 2005), except for two Sphagnum species in which they occupy
water-filled, dead (hyaline) cells, where they may gain protection from the low pH
of the bog environment (Solheim and Zielke 2002). The most common
cyanobacterial epiphytes on mosses are members of the filamentous, heterocystous
genera Nostoc, Stigonema and Calothrix (DeLuca et al. 2002, 2007; Gentili et al.
2005; Houle et al. 2006), although the non-heterocystous, filamentous genera
Phormidium and Oscillatoria, and even the unicellular Microcystis, have also
been reported (Solheim et al. 2004). These moss associations will not be discussed
further here as they are almost exclusively epiphytic, and even in the seemingly
endophytic Sphagnum symbioses the cyanobacteria occupy dead cells which are
connected to the outside environment. Nevertheless, these nitrogen-fixing
cyanobacteria–moss associations are of environmental significance as they are
often the major source of combined nitrogen in local ecosystems in the Arctic,
the Antarctic and forests of the northern hemisphere where mosses are abundant
(Zielke et al. 2002, 2005; Solheim and Zielke 2002; Nilsson and Wardle 2005;
DeLuca et al. 2008).
Fig. 1 The liverwort Blasia
pusilla collected from the
wild, showing the dark
Nostoc colonies
(~0.5–1.0 mm in diameter)
bordering the thallus midrib.
Reproduced with permission
from Adams (2000)
96
D.G. Adams and P.S. Duggan
Endophytic cyanobacterial symbioses are found in all 13 hornwort genera
(Renzaglia et al. 2007), but in only two (Blasia and Cavicularia) of the greater
than 340 liverwort genera, although a further two (Marchantia and Porella) form
epiphytic associations (Rai et al. 2000; Adams 2000, 2002a, b; Adams et al. 2006;
Adams and Duggan 2008). In the endophytic symbioses, cyanobacterial colonies
can be seen as dark spots up to 0.5 mm in diameter within the flattened thallus of the
plant (Fig. 1). The endophytes of liverworts and hornworts are almost exclusively
members of the genus Nostoc (Costa et al. 2001; Rasmussen and Nilsson 2002;
Adams 2002a, b; Adams et al. 2006; Bergman et al. 2007a; Adams and Duggan
2008), and a wide variety of strains can infect a single thallus in the field (West and
Adams 1997; Costa et al. 2001; Adams 2002a, b; Adams and Duggan 2008),
although some can be dominant and widespread (Rikkinen and Virtanen 2008).
The ease with which liverworts and hornworts can be grown in the laboratory in
shaken liquid culture (Fig. 2b), with or without symbiotic cyanobacteria, makes
them an excellent model for studying the infection process (Adams 2002a, b;
Meeks 2003; Duckett et al. 2004; Adams and Duggan 2008).
The structures housing the symbiotic cyanobacteria in hornworts (slime or
mucilage cavities; Fig. 2a) and liverworts (auricles; Fig. 2c, d) are produced
constitutively and not as a response to the presence of potential symbionts
(Renzaglia et al. 2000; Adams 2002a, b; Meeks 2003; Adams et al. 2006; Villarreal
and Renzaglia 2006; Bergman et al. 2008). The slime cavities of hornworts such as
Anthoceros and Phaeoceros are formed within the thallus and are connected to the
ventral surface by mucilage clefts (Fig. 2a) which superficially resemble stomata,
but are not thought to be related (Villarreal and Renzaglia 2006). The cleft results
from the separation of adjacent epidermal cells, after which the slime cavity
develops beneath the cleft (Renzaglia et al. 2000). The slime cavities in the
hornwort Leiosporoceros dussii also develop beneath mucilage clefts (Fig. 3c, d)
but are elongated mucilage-filled “canals” (Fig. 3b) resulting from the separation of
plant cell walls along their middle lamellae. These canals branch to form an
integrated network enabling the Nostoc symbiont to spread throughout the thallus
(Villarreal and Renzaglia 2006). By contrast, in the liverwort Blasia the
cyanobacteria occupy dome-shaped auricles (Fig. 2c, d), which develop on the
ventral surface of the thallus from a three-celled mucilage hair that undergoes
extensive elaboration (Renzaglia et al. 2000).
2.2
Gymnosperms (Cycads)
Cycads are evergreen palm-like plants, from 10 cm to 20 m in height, which once
dominated the Earth’s forests but are now restricted to subtropical and tropical
regions of mostly the southern hemisphere (Brenner et al. 2003; Vessey et al. 2005).
Cycads produce coralloid (coral-like) roots (Fig. 4a) that become infected with
heterocystous cyanobacteria (Costa and Lindblad 2002; Lindblad and Costa 2002;
Vessey et al. 2005; Lindblad 2009) usually from the genus Nostoc, although
Signalling in Cyanobacteria–Plant Symbioses
97
Fig. 2 The liverwort and hornwort symbioses. (a) Fluorescence micrograph of the hornwort
Phaeoceros sp. stained with calcofluor, showing the slit-like entrances (one of which is arrowed)
through which hormogonia gain entry to the slime cavities beneath. (b) View of the underside of an
Erlenmeyer flask containing the liverwort Blasia pusilla grown free of cyanobacteria in shaken
liquid medium. (c) Blasia pusilla growing in liquid culture showing three auricles infected in the
laboratory with two different Nostoc strains, one brown pigmented (the two auricles to the left) and
the other blue-green. (d) Fluorescence micrograph of uninfected Blasia stained with calcofluor. A
single auricle can be seen with one inner (lower arrow) and one outer (upper arrow) slime papilla.
Bars 50 mm. Photographs (a) and (d) courtesy of S. Babic. (a) and (d) reproduced with permission
from Adams (2000) (b) reproduced with permission from Adams (2002a) (c) reproduced with
permission from Adams and Duggan (1999)
Calothrix spp. have been reported on a number of occasions (Costa and Lindblad
2002; Rasmussen and Nilsson 2002; Bergman et al. 2007a, 2008; Gehringer et al.
2010; Thajuddin et al. 2010). Single or multiple strains can infect a coralloid root
(Zheng et al. 2002; Costa et al. 2004; Thajuddin et al. 2010) although some Nostoc
spp. can be dominant and, at least in the cycad genus Macrozamia, there seems to be
little host specialisation by cyanobionts in the field (Gehringer et al. 2010).
Coralloid roots grow out from the tap root, and then upwards, sometimes
breaking the soil surface. The mechanism by which the cyanobacteria enter the
roots is unknown, although suggestions include lenticels and breaks in the dermal
layer (Costa and Lindblad 2002; Vessey et al. 2005; Bergman et al. 2007a) possibly
resulting from degradation of the cell wall caused by bacteria and fungi in the cycad
98
D.G. Adams and P.S. Duggan
Fig. 3 The hornwort Leiosporoceros dussii with symbiotic Nostoc. (a) To the left is a young
rosette and to the right an older thallus with numerous upright sporophytes. (b) Dark green Nostoc
“strands” (arrows) can be seen within the thallus, parallel to the main axis. S sporophyte. (c) Light
micrograph of a nearly transverse section of two mucilage clefts (arrows), which provide the entry
point for cyanobacterial infection; the Nostoc filaments subsequently spread through channels
created by the separation of cells along their middle lamellae. (d) Scanning electron micrograph of
a mucilage cleft. Bars 10 mm in (a), 2 mm in (b), 15 mm in (c) and 20 mm in (d). Reproduced with
permission from Villarreal and Renzaglia (2006)
rhizosphere (Lobakova et al. 2003). The cyanobionts occupy a mucilage-filled zone
between the inner and outer cortical layers (Fig. 4b). Although coralloid roots
develop in the absence of cyanobacteria, infection induces morphological
alterations (Sect. 5.4) which increase the area of contact between plant cells and
the cyanobiont, to enhance nutrient exchange (Adams 2000; Rai et al. 2000; Costa
and Lindblad 2002; Lindblad 2009).
2.3
Angiosperms (Gunnera)
This is the only intracellular cyanobacterial-plant symbiosis (the Nostoc cells are
found between the host cell wall and the plasmalemma) and the only symbiosis with
a flowering plant (Bergman 2002; Bergman and Osborne 2002; Bergman et al.
2007a). The genus Gunnera consists of plants with leaves varying from 1 cm to
several metres across (Fig. 5a), once found only in warm, wet equatorial regions,
but which now commonly occur in suitably wet temperate regions (Osborne and
Sprent 2002). The cyanobiont is found in mucus-secreting stem glands (Figs. 5b
and 6a, b) and supplies the entire nitrogen requirements of even the largest plants
(Bergman 2002; Bergman et al. 2007a). In the field, Gunnera is infected by a wide
range of Nostoc spp. (Nilsson et al. 2000; Rasmussen and Svenning 2001; Guevara
et al. 2002; Svenning et al. 2005) all of which are capable of high levels of
Signalling in Cyanobacteria–Plant Symbioses
99
Fig. 4 The cycad–Nostoc
symbiosis. (a) Cycad
coralloid root, which is the
site of cyanobacterial
infection. (b) Transverse
section of the root showing
the dark cyanobacterial band
between the inner and outer
cortical layers of the root. (a)
Reproduced with permission
from Lindlbad et al. (1985)
(b) reproduced with
permission from
Rai et al. (2000)
differentiation into hormogonia (Bergman et al. 2007a, 2008). A single stem gland
may occasionally contain several different (but closely related) strains of Nostoc
(Nilsson et al. 2000). However, even closely related Nostoc strains can differ in
their ability to infect Gunnera (Papaefthimiou et al. 2008a) and some strains that
form large numbers of motile hormogonia are incapable of infecting (Nilsson et al.
2006), implying that the successful establishment of the symbiosis involves additional factors, perhaps including chemical signalling between the host and potential
cyanobionts (Sect. 4).
Cyanobacteria gain entry to Gunnera through the specialised stem glands found
at the base of each leaf stem (petiole; Fig. 6a, b). Mucilage secreted by these glands
contains unidentified chemical signals that both induce the formation of
hormogonia and attract them by chemotaxis (Bergman 2002; Bergman et al.
2007a). The formation of glands is not reliant on the presence of cyanobacteria,
although it is stimulated by nitrogen starvation (Chiu et al. 2005). The gland
consists of a central papilla surrounded by a further 5–8 papillae, creating narrow
channels that lead downwards into the stem tissues (Fig. 6b). Destruction of these
channels by removal of all of the outer papillae prevents cyanobacterial infection,
but this is restored by leaving just a single outer papilla (Uheda and Silvester 2001).
The channels therefore provide the route by which Nostoc hormogonia migrate
towards the plant tissue at the base of the gland, and once there, they enter the
Gunnera cells by an unknown mechanism; following cell invasion the host cell wall
appears normal (Bergman 2002; Bergman et al. 2007a). Symbiotically competent
100
D.G. Adams and P.S. Duggan
Fig. 5 Gunnera manicata. (a) A young plant with two large flower spikes. Inset: Red pigmented
fronds cover the crown of the plant (hidden in the large image), where new leaves and new stem
glands develop. (b) Vertical cross-section of a rhizome of Gunnera manicata. Cyanobacterial
colonies (0.5–2 cm in diameter) can be seen as green patches around the periphery of the rhizome.
New leaves will develop in the region between the two leaf petioles at the top of the image, which
is an area covered by red fronds (see inset in a). New stem glands form close to the base of each
newly developing leaf petiole and subsequently become infected by Nostoc. Photos: Owen
Jackson
Fig. 6 Gunnera stem glands. (a) Gunnera seedling showing the red stem glands at the base of the
leaf petioles. The glands are the entry point for cyanobacteria. (b) Scanning electron micrograph of
a Gunnera chilensis gland showing the arrangement of papillae. Hormogonia gain entry into the
internal stem gland tissue by migrating down the channels between the papillae. (a) reproduced
with permission from Adams et al. (2006) (b) reproduced with permission from Bergman et al.
(1992)
Signalling in Cyanobacteria–Plant Symbioses
101
Fig. 7 The water fern Azolla. (a) View from above, of Azolla filliculoides floating on the water
surface. (b) View of an Azolla branch showing the overlapping dorsal lobes of the leaves which
contain the cyanobionts. (c) Light micrograph of the Azolla cyanobiont with the large heterocysts
clearly visible. (d) Transmission electron micrograph of a thin, longitudinal section of a
cyanobiont filament showing a heterocyst (centre) with a vegetative cell on either side. (e)
Fluorescence micrograph of a pair of megasporocarps (blue) which become infected with
cyanobacteria when the motile hormogonia (h) on the surface, enter via channels (arrows).
Once inside the megasporocarp the cells of the hormogonia convert into a form of spore called
an akinete (ak) which can be seen as the intensely fluorescing area (red) above the megaspores
(sp). These akinetes provide the inoculum for the next generation of the fern, so maintaining the
continuity of the symbiosis. Bars 5 mm (c), 5 mm (d). Reproduced with permission from Ran et al.
(2010)
cyanobacteria are known to produce the phytohormone indole-3-acetic acid
(Sergeeva et al. 2002; Liaimer and Bergman 2004) which might be responsible
for localised mitotic activity of the host cells which are thought to be the site of
infection (Bergman et al. 2007a). The gland channels disappear at this stage,
preventing any further infection. The cyanobacteria fill the Gunnera cells but
never enter the cytoplasm as they are confined between the plasmalemma and the
cell wall, the former acting as the symbiotic interface where nutrients are
exchanged between the partners (Bergman 2002; Bergman et al. 2007a).
2.4
Pteridophytes (Azolla)
Water ferns of the genus Azolla have overlapping leaves consisting of two 1-mm
long lobes (Fig. 7a, b), the upper one containing diazotrophic cyanobacteria in an
ovoid cavity approximately 0.3 mm in length, and the lower achlorophyllous lobe
serving as a float to keep the upper lobe above the water surface (Adams 2000;
Lechno-Yossef and Nierzwicki-Bauer 2002; Carrapico 2002; van Hove and
Lejeune 2002a, b; Bergman et al. 2007a, b). These ferns are generally no larger
than 3–4 cm and are found on still or slow-moving freshwater bodies in temperate
102
D.G. Adams and P.S. Duggan
to tropical climates. The association has been used for centuries as a green manure
for rice cultivation (Vaishampayan et al. 2001; Bergman et al. 2007a, b) but also has
potential applications in mosquito control and as a supplemental animal feed (van
Hove and Lejeune 2002a, b; Choudhury and Kennedy 2004).
Azolla is unique among the cyanobacteria–plant symbioses in being the only
example in which the cyanobiont is perpetually associated with the host, being
transferred from generation to generation. Mature dorsal leaf cavities of Azolla
contain approximately 2,000–5,000 cyanobiont cells together with a complex
community of heterotrophic bacteria, all of which are immobilised within a polysaccharide-rich mucilage around the periphery of the cavity, the central region
being gas-filled (van Hove and Lejeune 2002a, b; Lechno-Yossef and NierzwickiBauer 2002). The wide range of bacteria found in the leaf cavities (and other parts
of the plant) are mostly unculturable (Bergman et al. 2007a; Zheng et al. 2009a, b).
Like the cyanobiont, these bacteria are present throughout the life cycle of the plant
and may therefore play an important role in the association, perhaps by contributing
to nitrogen fixation and the production of the polysaccharide-rich mucilaginous
matrix associated with the leaf cavities (Zheng et al. 2009a, b), or by releasing the
plant hormone auxin (Lechno-Yossef and Nierzwicki-Bauer 2002).
The Azolla leaf cavity is occupied by an abundant primary cyanobiont (Fig. 7c, d),
which appears to be unable to grow outside the symbiosis, and a much less abundant
secondary cyanobiont which can be readily cultured (Papaefthimiou et al. 2008a, b;
Sood et al. 2008a, b). These major and minor cyanobionts show significant phenotypic differences. In 1873, Strasburger was the first to refer to the major cyanobiont as
Nostoc, but this was later changed to Anabaena, since when there has been continuing
controversy about its correct generic assignment. The major cyanobiont is referred to
as Nostoc azollae 0708 in the recently published draft genome sequence (http://
genome.jgi-psf.org/anaaz/anaaz.home.html; Ran et al. 2010), although the closest
phylogenetic relatives appear to be Raphidiopsis brookii D9 and Cylindrospermopsis
raciborskii CS-505, which have the smallest sequenced genomes of the filamentous
cyanobacteria. However, Nostoc azollae 0708 shares the highest number of protein
groups exclusively with the symbiotically competent Nostoc punctiforme PCC 73102
(Ran et al. 2010). The primary cyanobiont has a small (5.49 Mb) genome, comprising
one chromosome and two plasmids, and contains 5,357 coding sequences of which
3,668 have intact open reading frames, while the rest are pseudogenes. High numbers
of pseudogenes are a trait associated with endosymbionts in sheltered environments
where the likelihood of encountering foreign DNA is low. The large proportion of
pseudogenes (31.2%) found within all genomic functions present in the genome of
the cyanobiont suggests a high level of gene erosion (Ran et al. 2010).
The permanent association between Azolla and its cyanobiont is maintained by
the transfer of the cyanobacteria to the progeny of both asexual and sexual reproduction. The former is the main form of reproduction and involves the doubling of
plant biomass in approximately 2 days, accompanied by fragmentation of branches
from the main stem. Cyanobionts, in the form of hormogonia-like, non-heterocystous filaments, are transferred to the cavities of newly developing leaves at the
apical meristem, with the help of bridge-like structures formed by primary branched
Signalling in Cyanobacteria–Plant Symbioses
103
hairs (Zheng et al. 2009a, b). Sexual reproduction of the fern seems to be triggered
by adverse environmental conditions (reviewed by Lechno-Yossef and NierzwickiBauer 2002; Pabby et al. 2004a) and involves sporophytes that produce male
microsporocarps and female megasporocarps, the latter containing a single megasporangium. Within the indusium chamber of each megasporangium is a single
megaspore and megaspore apparatus, together with a colony of the cyanobiont
which, in the form of hormogonia-like filaments, entered the chamber via a pore at
the top (Fig. 7e; Zheng et al. 2009b). The pore subsequently closes and the cells of
the filaments undergo synchronous conversion to spore-like resting cells known as
akinetes (Zheng et al. 2009b). In this way the mature, dormant megasporocarp
contains similarly dormant cyanobiont akinetes. Megasporocarp fertilisation and
embryogenesis are followed by akinete germination to form active cyanobiont
filaments that provide the inoculum for the embryonic leaf (Zheng et al. 2009a).
3 The Role of Hormogonia in Plant Infection
3.1
Hormogonia and Motility
The primary function of the cyanobiont in plant symbioses is to provide combined
nitrogen for the partnership and, accordingly, they are mostly heterocystous
cyanobacteria, primarily from the genus Nostoc. For cyanobacteria in the soil to
find a suitable plant host they must be motile, yet members of the genus Nostoc are
non-motile for most of their life cycle. However, they can produce short, motile
filaments known as hormogonia which serve both as a means of dispersal and as the
agent of plant infection (Meeks 2003, 2009; Adams 2000; Gusev et al. 2002; Meeks
et al. 2002; Meeks and Elhai 2002; Bergman et al. 2007a). Hormogonia development is triggered by a variety of environmental factors (Meeks et al. 2002; Meeks
and Elhai 2002; Meeks 2009) including chemicals released by plants (Sect. 4). The
process begins with a round of rapid, synchronous cell divisions in all vegetative
cells, followed by filament fragmentation at the junction between each heterocyst
and the neighbouring vegetative cell. This releases free heterocysts (which are no
longer viable) and short filaments which become the motile hormogonia. After a
period of dispersal the hormogonia lose motility and develop heterocysts once
more, enabling them to fix nitrogen; this is vital for a stable symbiosis and plants
secrete chemicals into the symbiotic tissues to prevent further hormogonia
formation.
Even the Azolla cyanobiont, which is retained by the fern from one generation to
the next, produces hormogonia-like filaments to ensure the transfer of the
cyanobiont to the megaspore during sexual reproduction, and to the new leaf cavities
that form following asexual reproduction (Sect. 2.4; Zheng et al. 2009a, b). However,
members of hormogonia-forming genera other than Nostoc are either rarely (e.g.
Stigonema and Calothrix) or never (e.g. Fischerella) found as plant symbionts,
104
D.G. Adams and P.S. Duggan
implying that additional factors are required for symbiotic competence. One such
factor is the ability to sense and react to plant compounds that act as
chemoattractants, guiding the hormogonia to the plant structures that will house
them. This is likely to be especially important in structures that receive no light,
because the natural phototaxis of hormogonia must be overridden to ensure successful infection. A second factor is the ability to adhere to and possibly to
specifically recognise the plant surface; for this, external filamentous protein
structures known as type IV pili are thought to be involved (see below). Finally,
plant cyanobionts need to be facultative heterotrophs to enable them to survive
in darkness in cycad roots and Gunnera stem glands where they can not
photosynthesise, but are provided with fixed carbon by the plant.
3.2
Pili
The surface of Nostoc hormogonia is covered by abundant type IV pili (Tfp; Fig. 8).
Tfp are found in a wide range of bacteria where they have roles in adhesion,
DNA uptake, pathogenesis and motility (Mattick 2002; Nudleman and Kaiser
2004; Burrows 2005). They are also involved in motility in some unicellular
cyanobacteria (Bhaya 2004), and their presence on the surface of motile
hormogonia, but absence from non-motile vegetative filaments, has led to the
suggestion that they are involved in the gliding motility of hormogonia (Duggan
et al. 2007). The N. punctiforme genome carries homologues of many of the genes
known to be involved in pilus biogenesis and function in other bacteria (Meeks
et al. 2001) and at least one of these, pilQ, is strongly upregulated during
hormogonia formation (Klint et al. 2006). In addition, mutation of pilT and pilD,
thought to be involved in Tfp function, greatly decreases the ability of the mutant
hormogonia to infect the liverwort Blasia (Duggan et al. 2007). However, it is not
clear if this is due to loss of motility (and hence, chemotaxis) or some other
potential function of the pili, such as adhesion to, or recognition of the plant surface
(Duggan et al. 2007).
3.3
Chemotaxis
The importance of signal sensing and chemotaxis in hormogonia is reflected in the
dramatic changes in gene expression that accompany hormogonia development
induced by either combined nitrogen starvation or hormogonia inducing factor
(HIF, Sect. 4; Campbell et al. 2007, 2008). In the first 24 h of N. punctiforme
hormogonia development, the transcription of 944 genes is upregulated and 856
downregulated (Campbell et al. 2007). A majority of the former encode proteins for
signal transduction and transcriptional regulation, with others encoding proteins
Signalling in Cyanobacteria–Plant Symbioses
105
Fig. 8 Comparison of the cell surfaces of wild-type vegetative filaments and hormogonia of the
symbiotic cyanobacterium Nostoc punctiforme. (a) Wild-type vegetative filaments; (b) wild-type
hormogonia. Scale bars represent 1 mm. For electron microscopy, platinum wire (2 cm by 0.2 mm)
was evaporated onto the surface of each sample by using an Edwards 306A high-vacuum coating
unit and samples were viewed on a JEOL1200EX transmission electron microscope at 80 kV.
Reproduced with permission from Duggan et al. (2007)
with putative roles in chemotaxis and pilus biogenesis (Meeks et al. 2001;
Klint et al. 2006; Campbell et al. 2007). Similarly, the genomes of symbiotic
cyanobacteria reflect the importance of signal sensing and transduction. For example, the N. punctiforme genome has 3–5 copies of genes encoding homologues of
chemotaxis-related proteins CheA, CheB, CheW, CheD and CheR (Meeks et al.
2001). Similarly, preliminary annotation of the draft genome sequence of the Azolla
filiculoides cyanobiont (http://genome.jgi-psf.org/anaaz/anaaz.home.html) has
revealed a large number of genes with potential involvement in signal perception
and transduction, in addition to genes encoding a possible pilus-related motility
apparatus.
3.4
Other Characteristics
In addition to the more obvious hormogonia traits of motility, chemotaxis and plant
recognition/adhesion, there is evidence that more subtle aspects of their behaviour
106
D.G. Adams and P.S. Duggan
are also crucial for successful plant infection. For example, in N. punctiforme
inactivation of the gene sigH, which encodes an alternative sigma subunit of
RNA polymerase, and transcription of which is induced by HIF (Meeks 2003),
has no effect on the frequency of HIF-induced hormogonia, yet the hormogonia are
fivefold more infective of the hornwort Anthoceros than are wild-type hormogonia
(Meeks and Elhai 2002; Meeks 2003); the reason for this is unknown.
The infection frequency of N. punctiforme hormogonia in the liverwort Blasia is
also influenced by mutations in cyaC, which encodes adenylate cyclase, the enzyme
responsible for the biosynthesis of adenosine 30 , 50 -cyclic monophosphate (cAMP),
implying that this intracellular messenger may play a role in infection (Adams and
Duggan 2008; Chapman et al. 2008). However, the CyaC adenylate cyclase is a
multidomain enzyme and mutations in different domains result in hormogonia with
wildly different infection frequencies in Blasia, with one showing a 300–400%
greater infection frequency than the wild type, and another showing only 25% of the
frequency of the wild type (Chapman et al. 2008). These different infection
phenotypes seem not to be the result of differences in cellular cAMP levels, as
these are 25% of wild type in both mutants. Nor do the mutants show any
differences in the frequency, motility or piliation of hormogonia induced in the
presence of Blasia. It seems, therefore, that differences in symbiotic competency
result from subtle, unknown shifts in the behavioural characteristics of the mutant
hormogonia in response to plant signals.
4 Signalling Between Potential Partners
Host plants increase their chances of infection by releasing compounds that trigger
hormogonia formation (hormogonia inducing factor, HIF) and others that act as
chemoattractants. Since hormogonia only remain infective while motile, plants
can further improve their chances of infection by prolonging the hormogonial
(motile) phase. For example, Gunnera stem gland mucilage contains a factor(s)
that lengthen this motile stage from 20–40 h to several weeks (see Bergman et al.
2007a). HIFs are produced by the hornwort Anthoceros punctatus (Meeks et al.
2002; Meeks and Elhai 2002; Meeks 2003, 2009), cycads and Gunnera (Rai
et al. 2000; Bergman et al. 2008) and the liverwort Blasia (Watts 2000; Adams
2002a, b; Adams and Duggan 2008), although none has been chemically identified
and they may differ in different plants. The production or release of HIF seems to be
stimulated by nitrogen starvation (Meeks and Elhai 2002; Meeks 2003, 2009). The
cyanobiont’s ability to respond to plant HIF influences the subsequent infection
frequency. For example, N. punctiforme mutants that show a greater hormogonia
induction in response to Anthoceros HIF also infect the plant at a greater initial
frequency than the wild type. Conversely, mutation of the N. punctiforme gene
ntcA, which encodes the global transcriptional regulator NtcA (Herrero et al. 2004),
reduces the frequency of HIF-induced hormogonia, and those that do form fail to
infect Anthoceros (Wong and Meeks 2002).
Signalling in Cyanobacteria–Plant Symbioses
107
Having stimulated hormogonia formation by the release of HIF a plant host must
then attract the hormogonia by the production of chemoattractants. A good example
is the liverwort Blasia, which releases a chemoattractant when nitrogen starved
(Knight and Adams 1996; Watts 2000; Adams and Duggan 2008). However, such
chemoattractants are not restricted to host plants as they are also produced by nonhost plants such as Trifolium repens (Nilsson et al. 2006) and germinating wheat
seeds (Knight and Adams 1996; Watts 2000; Adams and Duggan 2008). None of
these chemoattractants have been chemically identified although they are thought to
be sugar-based molecules (Watts 2000); indeed, simple sugars such as arabinose,
glucose and galactose have been shown to attract hormogonia, with arabinose being
the most effective (Nilsson et al. 2006).
The polysaccharide-rich mucilage secreted by mature stem glands of Gunnera
chilensis contains polymers of arabinose, galactose and glucuronic acid, with the
major form being arabinogalactans associated with arabinogalactan proteins (AGPs;
Rasmussen et al. 1996). AGPs are a diverse group of proteoglycans generally found
in the extracellular matrix of plant and some algal cells (Gaspar et al. 2001; Seifert
and Roberts 2007; Showalter 2001). Although AGPs are found in Gunnera stem
gland mucilage, their significance for the symbiosis is not known (see: Bergman
2002; Bergman et al. 2007a). However, they have been implicated in the
Alnus–Frankia symbiosis in which they are found at the symbiotic interface in the
root nodules (Berry et al. 2002). Gunnera mucilage contains high levels of arabinose, possibly released from AGPs or arabinan-containing pectins by the extracellular enzyme ARAf, expression levels of which are slightly higher in stem tissue
containing glands compared with that lacking glands (Khamar et al. 2010).
The high levels of sugars such as arabinose in Gunnera mucilage ensure
effective chemoattraction of hormogonia into the gland channels, and from there
into the deeper tissues. This is aided by the very low levels of the reducing sugars
glucose, fructose and sucrose, which are known to suppress hormogonia formation
(Khamar et al. 2010). However, these sugars are found at 100-fold higher
concentrations in the mature gland tissue from which the mucilage originates,
ensuring that the development of hormogonia that were attracted by arabinose in
the mucilage and gland channels, is suppressed in mature gland tissue, facilitating
heterocyst development and nitrogen fixation. The higher levels of glucose and
fructose compared with sucrose in mature gland tissue may result from the high
expression levels of genes encoding sucrose-hydrolyzing enzymes, such as cell wall
invertase, in stem tissue containing glands compared with tissue lacking glands
(Khamar et al. 2010). Although starch is abundant in the cortical cells of stems of
nitrogen-starved Gunnera seedlings, mature gland tissue is devoid of starch, perhaps as a result of the increased expression of genes encoding enzymes for starch
degradation, including starch phosphorylase and a-amylase (Khamar et al. 2010).
In the Gunnera symbiosis, stem gland mucilage contains a compound(s) that
stimulates expression of Nostoc genes hiaA, hiaB and hiaC which encode, respectively, an outer membrane glycoprotein, a putative signalling molecule and a
protein that may be involved in the adaptation of Nostoc to an acidic environment
such as that in stem gland mucilage (Liaimer et al. 2001; Bergman et al. 2007a).
108
D.G. Adams and P.S. Duggan
However, as might be expected, the expression of many more Nostoc genes is
altered in planta. For example, proteomic analysis has identified 38 Nostoc proteins
differentially expressed in symbiosis, including four upregulated cell surfaceassociated proteins one of which contains fasciclin-like repeats (Ekman et al.
2006). Such fasciclin domains are typical of surface-associated proteins involved
in cell adhesion, which are thought to be symbiotically relevant in several unrelated
symbioses including Nostoc-containing lichens (Paulsrud and Lindblad 2002).
Another group of signalling compounds with potential involvement in the
cyanobacterial symbioses is the lectins. These are produced by the plant host in
bryophyte and Azolla symbioses, and can bind to sugars on the surface of symbiotic
Nostoc strains (Lehr et al. 2000; see also: Rai et al. 2000; Adams 2000; Rikkinen
2002; Adams et al. 2006). Little is known about the importance of lectins in
cyanobacteria–plant symbioses, although they have been suggested to be involved
in fungus-partner recognition in lichens (Lehr et al. 2000; Elifio et al. 2000;
Rikkinen 2002; Legaz et al. 2004; Sacristan et al. 2006). The majority of lichens
consist of a fungus and a green alga, but around 10% contain a cyanobacterial
partner either as sole photobiont, or in combination with a green alga (Rikkinen
2002; Adams 2011). In the cyanolichens Peltigera canina and Leptogium
corniculatum, a fungal arginase acts as a lectin by binding to a polygalactosylated
urease in the cell wall of the Nostoc cyanobiont (Diaz et al. 2009; Vivas et al. 2010).
Lectin binding may also be important in the unique cyanobacteria–fungus association G. pyriformis (Adams et al. 2006), as mannose-specific lectin ConA shows
changes in its binding during the Nostoc life cycle, with strong binding only to socalled primordia, which are hormogonia that have lost motility and are beginning to
develop heterocysts (Kluge et al. 2002; Adams et al. 2006). Primordia are also the
only stage which can be engulfed by the fungus to establish the symbiosis, implying
that Nostoc extracellular glycoconjugates could be important in recognition.
5 Host–Cyanobiont Interactions Post-infection
Cyanobacteria in symbiosis with plants generally show morphological
modifications such as the cell enlargement and shape irregularity seen in hornwort-associated Nostoc (Meeks and Elhai 2002) and in the cyanobiont of Azolla
(Pabby et al. 2003, 2004b; Papaefthimiou et al. 2008a; Sood et al. 2008a, b; Zheng
et al. 2009a). Further changes include repression of hormogonia development,
reduction of cell division, increases in heterocyst frequency and nitrogen fixation,
and repression of nitrogen assimilation and photosynthetic CO2 fixation. Such
changes are often least pronounced in the youngest tissue (the most recently
infected) and greatest in the oldest symbiotic tissues.
Signalling in Cyanobacteria–Plant Symbioses
5.1
109
Repression of Hormogonia Development
Although hormogonia are essential for a cyanobiont to locate and invade a plant host,
they lack heterocysts and are therefore incapable of nitrogen fixation, so their
formation must be repressed once the cyanobiont is inside the host. In the hornwort
A. punctatus, the plant achieves this repression by releasing into the symbiotic cavity
an unidentified water-soluble hormogonia-repressing factor (HRF) which is dominant
over HIF (Cohen and Yamasaki 2000; Meeks and Elhai 2002; Meeks et al. 2002;
Campbell et al. 2003; Meeks 2003). HRF induces expression of the N. punctiforme
gene hrmA and strains mutated in this gene form hormogonia in the presence of HIF.
hrmA is part of the hrmRIUA operon which resembles sugar uronate metabolism
operons of other bacteria, although hrmA itself has no sequence homology with any
gene in the databases (Campbell et al. 2003; Meeks 2003). The hrm locus is central to
repressing further hormogonia formation in the Anthoceros–Nostoc symbiosis. The
gene product of hrmR is a transcriptional repressor belonging to the LacI/GalR
family of sugar-binding repressors and binds galacturonate in vitro (Campbell et al.
2003). The gene is self-regulating and also regulates hrmE, whose function is
unknown. Both hrmR and hrmE are negatively regulated by fructose (Ungerer
et al. 2008). Interestingly, immediately downstream of hrmE are homologues of
genes involved in fructose transport (Ungerer et al. 2008) that are known to be
induced by HRF and are therefore considered to be part of the hrm locus (Meeks,
2006). Hormogonia formation is known to be repressed by specific sugars, with
sucrose being the most effective, followed by glucose and fructose (Khamar et al.
2010). Taken together these observations raise the possibility that fructose (possibly
converted to a signalling metabolite) might regulate hormogonia formation
(Ungerer et al. 2008). Indeed, the high concentrations of glucose and fructose in
Gunnera manicata mature stem gland tissue led Khamar et al. (2010) to apply the
model of Campbell et al. (2003) and conclude that hrmR would be sugar bound and
therefore inactive, thus ensuring the expression of genes that negatively regulate
hormogonia formation.
In summary, it would appear that free arabinose and galactose present in the
Gunnera mucilage efficiently attract Nostoc hormogonia (Sect. 4; Nilsson et al.
2006) but do not interfere with their formation, allowing them to reach their sites of
infection within the host plant. However, inside the Gunnera stem gland tissue the
high levels of glucose and fructose suppress further hormogonia formation and
promote heterocyst differentiation and nitrogen fixation (Khamar et al. 2010).
The hrm operon may be involved in symbiotic systems other than Anthoceros
because expression of hrmA in N. punctiforme is induced by the plant flavonoid
naringin (Cohen and Yamasaki 2000) and by aqueous extracts of fronds of Azolla
pinnata and A. filiculoides (Cohen et al. 2002). Expression of hrmA is also induced
by deoxyanthocyanin (the pigment that gives Azolla its reddish colour in winter) in
synergy with other plant-derived compounds (Cohen et al. 2002). The pigment may
be a component of Azolla HRF as it is more abundant in mature frond tissue than in
110
D.G. Adams and P.S. Duggan
actively growing apical regions where hormogonia-like filaments are involved in
the infection of newly formed leaf cavities (Cohen et al. 2002).
5.2
Cell Division Control
The cyanobionts of plants have the potential for much more rapid growth than the
host and so in symbiosis their growth must be controlled to ensure the stability of
the partnership. Although not well understood, this growth control is thought to be
achieved by a variety of means such as the blocking of cell division, the restriction
of the number and size of symbiotic structures (by, for example, physical confinement) and restriction of the nutrient supply (Rai et al. 2000; Ekman et al. 2006;
Bergman et al. 2007a). For example, the Nostoc cyanobiont of the hornwort
Anthoceros grows up to tenfold more slowly in symbiosis than when free living
(Meeks 2003). Anthoceros also regulates the biomass and rate of nitrogen fixation
of its Nostoc colonies to maintain a stable rate of nitrogen fixation per unit of plant
tissue when, for example, growth is stimulated by elevated CO2 or light (Meeks and
Elhai 2002; Meeks 2003). The mechanism by which this is achieved is not known.
In Azolla the growth rates of both host and cyanobiont follow a decreasing linear
gradient from their maximum in new leaves at the apical end of the fronds, to a
minimum in the older leaves in the mature regions of the plant (Bergman et al.
2007a, b). The host appears to directly regulate cyanobiont growth because when
plant growth is artificially reduced with the inhibitor of eukaryotic protein synthesis, cycloheximide, cell division of the cyanobiont in leaves at the plant apex also
stops. Cyanobiont cell size and numbers per leaf cavity increase from apical to
older regions of the fern (leaf numbers 1–15), after which (leaves 15–28) the
numbers per cavity become constant, whereas cell size continues to increase
(Bergman et al. 2007a, b).
5.3
Heterocyst Development
Although the presence of heterocysts is essential for a stable nitrogen-fixing
symbiosis, mutants unable to differentiate heterocysts may still be able to infect a
host plant even though they are unable to support growth of the plant in the absence
of combined nitrogen. This is true for mutants of N. punctiforme inactivated in hetR
and hetF (Wong and Meeks 2002); the former gene encodes the primary driver of
heterocyst development (Golden and Yoon 2003; Zhang et al. 2006), and the gene
product of the latter is involved in the regulation and localisation of hetR transcription (Wong and Meeks 2001). Although incapable of fixing nitrogen for the
symbiosis both mutants infect the hornwort Anthoceros as efficiently as the wild
type. By contrast, inactivation of the ntcA gene, encoding the global nitrogen
Signalling in Cyanobacteria–Plant Symbioses
111
regulator NtcA (Flores and Herrero 2005), completely prevents infection even
though the mutant makes motile hormogonia (Wong and Meeks 2002).
Plant hosts are capable of supplying fixed carbon for both themselves and the
cyanobiont, enabling the latter to increase nitrogen fixation capacity by committing
a greater number of CO2-fixing vegetative cells to become heterocysts (which do
not fix CO2). By adopting a heterotrophic mode of nutrition the cyanobionts can
also grow in the roots of cycads and the stem glands of Gunnera, where there is
little or no light. The cyanobionts in plants therefore generally have heterocyst
frequencies greatly elevated above the 5% typical of free-living strains. However,
heterocyst frequency is not uniform throughout the plant, being lowest (typically
10–15%) in rapidly growing regions (such as the root tip of cycads), increasing to
60% or more in the oldest symbiotic tissue where multiple adjacent heterocysts are
often found, although these are very rare in free-living cyanobacteria. In these
regions, some of the heterocysts are likely to be senescent or dead (Meeks and Elhai
2002; Meeks 2003, 2009) and, perhaps as a result of this, the highest nitrogen
fixation rates are found at intermediate heterocyst frequencies; this is also the
region of the Gunnera stem where cyanobiont hetR expression is greatest (Wang
et al. 2004).
An important question is what regulates heterocyst frequency in planta? There is
some evidence that the environmental conditions experienced by the cyanobionts,
such as low light and a high concentration of host-derived sugars, are responsible
for the elevated heterocyst frequencies. For example, the growth of the Gunnera
isolate Nostoc PCC 9229 in the dark in the presence of fructose results in the
formation of double and quadruple heterocysts, which are not seen in the absence of
the sugar (Wouters et al. 2000).
In free-living cyanobacteria heterocyst development is triggered by nitrogen
starvation, which is thought to be perceived via an increase in the intracellular
concentration of 2-oxoglutarate (Muro-Pastor et al. 2001; Vazquez-Bermudez et al.
2002; Zhang et al. 2006) which activates the transcriptional regulator NtcA
(Herrero et al. 2004; Flores and Herrero 2005; Muro-Pastor et al. 2005). In turn,
NtcA enhances transcription of genes encoding both positive (e.g. HetR and HetF)
and negative (e.g. PatN and PatS) regulators of heterocyst development (Meeks and
Elhai 2002; Herrero et al. 2004; Zhang et al. 2006; Meeks 2009). However,
symbiotically associated cyanobacteria do not show the physiological characteristics of nitrogen limitation (Sect. 6) implying that the signal for heterocyst development in symbiosis is supplied by the host. A similar conclusion can be drawn
from observations of the behaviour of a N. punctiforme mutant defective in the
assimilation of nitrate (Meeks and Elhai 2002; Meeks 2003), which imply that in
planta the regulation of nitrogen fixation and heterocyst development is plant
mediated and independent of the nitrogen status of the cyanobiont (Meeks 2003,
2009). The identity of this plant signal and its target(s) are unknown but it is likely
to act prior to activation of the key heterocyst differentiation gene hetR (Zhang et al.
2006) and also prior to ntcA (discussed further by Meeks and Elhai 2002; Wong and
Meeks 2002; Meeks 2009).
112
5.4
D.G. Adams and P.S. Duggan
Host Changes
Morphological changes are apparent in both host and cyanobiont following infection, although the most dramatic changes are found in the latter. Such changes in the
host often reflect the need for efficient nutrient exchange and they result in
increased physical intimacy of the partners. This is not necessary in Gunnera
because the intracellular location of the Nostoc ensures efficient transfer of
nutrients. However, the area of contact between host and cyanobiont is increased
in some bryophytes by the production of multicellular filaments that grow from the
auricle wall in the liverwort Blasia and the wall of the slime cavity in the hornwort
A. punctatus, and infiltrate the cyanobacterial colony (see: Adams 2002a, b; Adams
and Duggan 2008). However, such wall ingrowths are absent in many other
hornworts, including Leiosporoceros (Villarreal and Renzaglia 2006). In cycads,
the area of cyanobiont–host contact is increased by elongation of the cells linking
the root inner and outer cortical layers (Costa and Lindblad 2002), creating a space
filled with mucilage and tightly packed cyanobacterial filaments and traversed by
specially elongated host cells (Vessey et al. 2005).
6 N2 Fixation and Transfer of Fixed Nitrogen
A significant proportion of the nitrogen fixed by plant cyanobionts is released to the
host (Table 1); this varies from 50% in Azolla to as much as 80–90% in Gunnera
and hornworts (Adams 2000; Meeks and Elhai 2002; Adams 2002a, b; Meeks 2003,
2009; Bergman et al. 2007a). In liverworts, hornworts and Azolla nitrogen is
released in the form of ammonia (Meeks 2003; Bergman et al. 2007a), whereas in
Gunnera it is ammonia and some asparagine (Bergman 2002; Bergman et al.
2007a), and in cycads it is thought to be glutamine and/or citrulline, depending
on the cycad genus (Table 1; Costa and Lindblad 2002; Vessey et al. 2005;
Lindblad 2009).
Cyanobacteria assimilate ammonia via the glutamine synthetase-glutamate
synthase (GS-GOGAT) pathway (Muro-Pastor et al. 2005; Flores and Herrero
2005) and ammonia release is mostly (with the exception of cycad cyanobionts) a
consequence of decreased cyanobiont GS activity resulting from either a reduction
in the specific activity of the enzyme or in the amount of protein produced,
depending on the host (Table 1). In hornworts, the level of GS protein is similar
in free living and symbiotically associated Nostoc implying that the observed
decrease in overall GS activity in symbiosis is due to a post-translational modification of the protein (Meeks and Elhai 2002; Meeks 2003, 2009).
Because cyanobionts release a large proportion of the nitrogen they fix, they
might be expected to show the signs of nitrogen starvation (Meeks and Elhai 2002).
In free-living cyanobacteria, this would be evident by degradation of the specialised
nitrogen storage compound cyanophycin (a co-polymer of arginine and aspartic
Signalling in Cyanobacteria–Plant Symbioses
113
Table 1 Heterocyst frequency and the characteristics of glutamine synthetase in symbiotically
associated cyanobacteria
Glutamine synthetaseb
Form of combined nitrogen
Host
Heterocyst
a
released (% released)c
frequency (%)
Amount of
Specific
protein (%)
activity (%)
Cycads
17–46
100
100
Glutamine/citrulline? (ND)
Gunnera
20–60
100
70
NH4+ (90)
Hornworts and 25–45
86–100
~15
NH4+ (80)
liverworts
Azolla
26–45
5–40
~30
NH4+ (40)
Lichens
Bipartite
4–8
<10
<10
NH4+ (90)
Tripartite
10–55
<10
<10
NH4+ (90)
ND not determined
Table compiled from Meeks (2009) and Adams (2000) and references therein
a
Heterocyst frequency is expressed as a percentage of the sum of heterocysts plus vegetative cells.
The range of frequencies reflects the increasing heterocyst frequency found with increasing age of
symbiotic tissue. Typical frequencies for free-living cyanobacteria are 4–10%
b
Glutamine synthetase is expressed as the amount of protein or the specific activity, both expressed
as a percentage of the value in the same cyanobacterium growing in the free-living state
c
The form of combined nitrogen released by the cyanobacterium to the host is given, with (in
parenthesis) the amount of nitrogen released by the cyanobiont as a percentage of the total nitrogen
fixed
acid) and proteins in carboxysomes (Rubisco) and in phycobilisomes (containing
accessory photopigments, the phycobiliproteins). However, all of these reserves are
found in relative abundance in symbiotically associated cyanobacteria.
7 CO2 Assimilation and Transfer of Carbon
Plant cyanobionts receive much of their fixed carbon from the host and their own
CO2 fixation capacity is greatly reduced (Table 2) as they adapt to a photo- or
chemo-heterotrophic form of nutrition (Meeks and Elhai 2002; Adams 2002a;
Meeks 2003). In the hornwort Anthoceros, this seems to be achieved by an
unknown post-translational modification of the enzyme Rubisco (ribulose-1, 5bisphosphate carboxylase/oxygenase, the primary carboxylating enzyme in
cyanobacteria), as the amount of the protein differs little between free-living or
Anthoceros-associated Nostoc (Meeks and Elhai 2002; Meeks 2003). However, the
mechanism must differ in the cyanobionts of cycads (Rai et al. 2000; Adams 2000;
Costa and Lindblad 2002) as Rubisco is active in extracts of freshly isolated
cyanobiont (Table 2), despite their heterotrophic mode of nutrition in planta
(Lindblad 2009). Photosynthetic CO2 fixation seems to be reduced in the Nostoc
symbiont of Gunnera by modifications of key photosynthetic components (such as
the D1 protein of the photosystem II complex; Black and Osborne 2004). In Azolla
Rubisco mRNA is 5–7 fold less abundant in the cyanobiont compared with free-
114
D.G. Adams and P.S. Duggan
Table 2 Characteristics of light-dependent CO2 fixation and ribulose bisphosphate carboxylase in
symbiotically associated cyanobacteria
Rubisco
Host
Light-dependent
CO2 fixation (%) a Protein (%)a Specific activity (%)a
Cycads
0
ND
87–100
Gunnera
<2
100
100
Hornwort (Anthoceros punctatus) 12
100
12–15
Azolla
85–100
ND
ND
Lichens
Bipartite
ND
~100
ND
Tripartite
~8
~100
~8
ND not determined
Table compiled from Meeks (2009) and Adams (2000) and references therein
a
Percentages given are the value in the cyanobiont immediately after isolation from the host,
compared with that in the free-living strain
living cultures (Adams 2000; Rai et al. 2000; Meeks 2009), yet immediately after
isolation from the plant the primary cyanobiont has approximately 85% of the
photosynthetic rate of free-living cyanobacteria (Table 2); why this should be is not
known.
8 Artificial Cyanobacteria–Plant Symbioses
The capacity of cyanobacteria for nitrogen fixation has clear potential applications
in agriculture for the supply of combined nitrogen to plants, but there are no
cyanobacterial endosymbioses with crop plants. Only Azolla has been used as a
green manure in agriculture, being grown in rice fields, releasing its nitrogen to the
soil upon death and decay. However, attempts to induce novel interactions between
cyanobacteria and plants have been numerous. Cyanobacteria have been introduced
into higher plant protoplasts (see: Rai et al. 2000; Adams 2000; Gusev et al. 2002;
Bergman et al. 2007a), or co-cultured with plant tissue cultures, plant regenerates
and cuttings from a variety of plants (Gantar 2000b; Gorelova 2001, 2006;
Lobakova et al. 2001a, b; Gorelova and Korzhenevskaya 2002; Gorelova and
Kleimenov 2003; Gorelova and Baulina 2009; see also Rai et al. 2000 and Gusev
et al. 2002 for a discussion of the earlier literature).
Some Nostoc hormogonia can be attracted by exudates of non-host plants
(Nilsson et al. 2006) and cyanobacteria can also colonise the surface of the roots
of rice (Nilsson et al. 2002; 2005) and wheat (Karthikeyan et al. 2007, 2009).
Colonisation of wheat seedling roots can enhance plant nitrogen content and root
growth (for a discussion of this, see Rai et al. 2000; Adams 2000; Gusev et al. 2002;
Bergman et al. 2007a) and cyanobacteria can be induced to grow within root tissues
by mechanical damage of the root (Gantar 2000a). Little is known about the extent
of such interactions in the field, but the enhancement of such “natural” interactions
Signalling in Cyanobacteria–Plant Symbioses
115
may offer the best hope of employing cyanobacteria to enhance crop growth and
replace at least a proportion of the current artificial fertiliser use. This might
involve, for example, producing crop plants with enhanced release of factors that
stimulate hormogonia development and chemoattraction, thus increasing root
colonisation. Certainly, this would be a great deal less technically challenging
than the creation of novel endosymbiotic associations with crop plants, in which
the cyanobiont is retained between host generations. Indeed, in the long evolutionary history of cyanobacteria–plant symbioses Azolla is the only such symbiosis to
have evolved.
Because most research to date has focussed on the cyanobacteria, our understanding of the plant host involvement in these symbioses is relatively poor. The recent
work of Khamar et al. (2010) examining changes in the expression of Gunnera genes
encoding enzymes for starch and sucrose hydrolysis is the first real attempt to assess
changes in host gene expression in response to the symbiotic state. Nevertheless, we
still know relatively little about the chemical signals and molecular mechanisms
involved in the establishment of a stable cyanobacteria–plant symbiosis.
Acknowledgements Our thanks go to the authors who gave permission for us to use the images
reproduced in this chapter.
References
Adams DG (2000) Symbiotic interactions. In: Whitton B, Potts M (eds) Ecology of cyanobacteria:
their diversity in time and space. Kluwer, Dordrecht, pp 523–561
Adams DG (2002a) Cyanobacteria in symbiosis with hornworts and liverworts. In: Rai AN,
Bergman B, Rasmussen U (eds) Cyanobacteria in symbiosis. Kluwer, Dordrecht, pp 117–135
Adams DG (2002b) The liverwort-cyanobacterial symbiosis. Biol Environ Proc R Ir Acad
102B:27–30
Adams DG (2011) Cyanobacterial symbioses. In: Whitton B (ed.) Ecology of cyanobacteria II.
Springer, Heidelberg
Adams DG, Duggan PS (1999) Heterocyst and akinete differentiation in cyanobacteria. New
Phytol 144:3–33
Adams DG, Duggan PS (2008) Cyanobacteria-bryophyte symbioses. J Exp Bot 59:1047–1058
Adams DG, Bergman B, Nierzwicki-Bauer SA, Rai AN, Schussler A (2006) Cyanobacterial-plant
symbioses. In: Dworkin M, Falkow S, Rosenberg E, Schleifer K-H, Stackebrandt E (eds) The
prokaryotes. a handbook on the biology of bacteria, vol 1, 3rd edn, Symbiotic associations,
biotechnology, applied microbiology. Springer, New York, pp 331–363
Bergman B (2002) The Nostoc-Gunnera symbiosis. In: Rai AN, Bergman B, Rasmussen U (eds)
Cyanobacteria in symbiosis. Kluwer, Dordrecht, pp 207–232
Bergman B, Osborne B (2002) The Gunnera-Nostoc symbiosis. Biol Environ Proc R Ir Acad
102B:35–39
Bergman B, Johansson C, S€
oderb€ack E (1992) The Nostoc-Gunnera symbiosis. New Phytol
122:379–400
Bergman B, Rasmussen U, Rai AN (2007a) Cyanobacterial associations. In: Elmerich C, Newton WE
(eds) Associative and endophytic nitrogen-fixing bacteria and cyanobacterial associations.
Kluwer, Dordrecht, pp 257–301
116
D.G. Adams and P.S. Duggan
Bergman B, Zheng W, Ekman M, Ran L (2007b) The cyanobacterium-Azolla symbiosis:
interactions and cell differentiation. Comp Biochem Physiol A Mol Integr Physiol 146A:S218
Bergman B, Ran L, Adams DG (2008) Cyanobacterial-plant symbioses: signaling and development. In: Herrero A, Flores E (eds) The cyanobacteria: molecular biology, genomics and
evolution. Caister, Norfolk, pp 447–473
Berry AM, Rasmussen U, Bateman K, Huss-Danell K, Lindwall S, Bergman B (2002)
Arabinogalactan proteins are expressed at the symbiotic interface in root nodules of Alnus
spp. New Phytol 155:469–479
Bhaya D (2004) Light matters: phototaxis and signal transduction in unicellular cyanobacteria.
Mol Microbiol 53:745–754
Black K, Osborne B (2004) An assessment of photosynthetic downregulation in cyanobacteria
from the Gunnera-Nostoc symbiosis. New Phytol 162:125–132
Brenner ED, Stevenson DW, Twigg RW (2003) Cycads: evolutionary innovations and the role of
plant-derived neurotoxins. Trends Plant Sci 8:446–452
Burrows LL (2005) Weapons of mass retraction. Mol Microbiol 57:878–888
Campbell EL, Wong FCY, Meeks JC (2003) DNA binding properties of the HrmR protein of
Nostoc punctiforme responsible for transcriptional regulation of genes involved in the differentiation of hormogonia. Mol Microbiol 47:573–582
Campbell EL, Summers ML, Christman H, Martin ME, Meeks JC (2007) Global gene expression
patterns of Nostoc punctiforme in steady state dinitrogen-grown heterocyst-containing
cultures, and at single time points during the differentiation of akinetes and hormogonia.
J Bacteriol 189:5247–5256
Campbell EL, Christman H, Meeks JC (2008) DNA microarray comparisons of plant factor- and
nitrogen deprivation-induced hormogonia reveal decision-making transcriptional regulation
patterns in Nostoc punctiforme. J Bacteriol 190:7382–7391
Carrapico F (2002) The Azolla-Anabaena-bacteria system as a natural microcosm. In: Hoover RB,
Levin GV, Paepe RR, Rozanov AY (eds) Instruments, methods, and missions for astrobiology
IV, Proceedings of SPIE vol 4495, pp 261–265
Chapman KE, Duggan PS, Billington NA, Adams DG (2008) Mutation at different sites in the
Nostoc punctiforme cyaC gene, encoding the multiple-domain enzyme adenylate cyclase, results
in different levels of infection of the host plant Blasia pusilla. J Bacteriol 190:1843–1847
Chiu WL, Peters GA, Levieille G, Still PC, Cousins S, Osborne B, Elhai J (2005) Nitrogen
deprivation stimulates symbiotic gland development in Gunnera manicata. Plant Physiol
139:224–230
Choudhury A, Kennedy IR (2004) Prospects and potentials for systems of biological nitrogen
fixation in sustainable rice production. Biol Fertil Soils 39:219–227
Cohen MF, Yamasaki H (2000) Flavonoid-induced expression of a symbiosis-related gene in the
cyanobacterium Nostoc punctiforme. J Bacteriol 182:4644–4646
Cohen MF, Sakihama Y, Takagi YC, Ichiba T, Yamasaki H (2002) Synergistic effect of
deoxyanthocyanins from symbiotic fern Azolla spp. on hrmA gene induction in the cyanobacterium Nostoc punctiforme. Mol Plant Microbe Interact 15:875–882
Costa JL, Lindblad P (2002) Cyanobacteria in symbiosis with cycads. In: Rai AN, Bergman B,
Rasmussen U (eds) Cyanobacteria in symbiosis. Kluwer, Dordrecht, pp 195–205
Costa JL, Paulsrud P, Rikkinen J, Lindblad P (2001) Genetic diversity of Nostoc symbionts
endophytically associated with two bryophyte species. Appl Environ Microbiol 67:4393–4396
Costa JL, Romero EM, Lindblad P (2004) Sequence based data supports a single Nostoc strain in
individual coralloid roots of cycads. FEMS Microbiol Ecol 49:481–487
DeLuca TH, Zackrisson O, Nilsson MC, Sellstedt A (2002) Quantifying nitrogen-fixation in
feather moss carpets of boreal forests. Nature 419:917–920
DeLuca TH, Zackrisson O, Gentili F, Sellstedt A, Nilsson MC (2007) Ecosystem controls on
nitrogen fixation in boreal feather moss communities. Oecologia 152:121–130
DeLuca TH, Zackrisson O, Gundale MJ, Nilsson M-C (2008) Ecosystem feedbacks and nitrogen
fixation in boreal forests. Science 320:1181
Signalling in Cyanobacteria–Plant Symbioses
117
Diaz E-M, Sacristan M, Legaz M-E, Vicente C (2009) Isolation and characterization of a
cyanobacterium-binding protein and its cell wall receptor in the lichen Peltigera canina.
Plant Signal Behav 4:598–603
Duckett JG, Burch J, Fletcher PW, Matcham HW, Read DJ, Russell AJ, Pressel S (2004) In vitro
cultivation of bryophytes: a review of practicalities, problems, progress and promise. J Bryol
26:3–20
Duggan PS, Gottardello P, Adams DG (2007) Molecular analysis of genes in Nostoc punctiforme
involved in pilus biogenesis and plant infection. J Bacteriol 189:4547–4551
Ekman M, Tollback P, Klint J, Bergman B (2006) Protein expression profiles in an endosymbiotic
cyanobacterium revealed by a proteomic approach. Mol Plant Microbe Interact 19:1251–1261
Elifio SL, Da Silva M, Iacomini M, Gorin PAJ (2000) A lectin from the lichenized Basidiomycete
Dictyonema glabratum. New Phytol 148:327–334
Flores E, Herrero A (2005) Nitrogen assimilation and nitrogen control in cyanobacteria. Biochem
Soc Trans 33:164–167
Flores E, Herrero A (2010) Compartmentalised function through cell differentiation in filamentous
cyanobacteria. Nat Rev Microbiol 8:39–50
Gantar M (2000a) Mechanical damage of roots provides enhanced colonization of the wheat
endorhizosphere by the dinitrogen-fixing cyanobacterium Nostoc sp. strain 2S9B. Biol Fertil
Soils 32:250–255
Gantar M (2000b) Co-cultivation of N2-fixing cyanobacterium Nostoc sp strain 2S9B and wheat
callus. Symbiosis 29:1–18
Gaspar Y, Johnson KL, McKenna JA, Bacic A, Schultz CJ (2001) The complex structures of
arabinogalactan proteins and the journey towards understanding function. Plant Mol Biol
47:161–176
Gehringer MM, Pengelly JJL, Cuddy WS, Fieker C, Forster PI, Neilan BA (2010) Host selection of
symbiotic cyanobacteria in 31 species of the Australian cycad genus: Macrozamia
(Zamiaceae). Mol Plant Microbe Interact 23:811–822
Gentili F, Nilsson MC, Zackrisson O, DeLuca TH, Sellstedt A (2005) Physiological and molecular
diversity of feather moss associative N2-fixing cyanobacteria. J Exp Bot 56:3121–3127
Golden JW, Yoon HS (2003) Heterocyst development in Anabaena. Curr Opin Microbiol
6:557–563
Gorelova OA (2001) Surface ultrastructure of the heteromorphic cells of Nostoc muscorum CALU
304 in a mixed culture with the Rauwolfia callus tissue. Microbiology 70:285–294
Gorelova OA (2006) Communication of cyanobacteria with plant partners during association
formation. Microbiology 75:465–469
Gorelova OA, Baulina OI (2009) Ultrastructure of cyanobacterium Nostoc sp f. Blasia cell forms
in persisting populations. Microbiology 78:609–617
Gorelova OA, Kleimenov SY (2003) The accumulation and degradation dynamics of cyanophycin
in cyanobacterial cells grown in symbiotic associations with plant tissues and cells. Microbiol
72:318–326
Gorelova OA, Korzhenevskaya TG (2002) Formation of giant and ultramicroscopic forms of
Nostoc muscorum CALU 304 during cocultivation with Rauwolfia tissues. Microbiol
71:563–569
Guevara R, Armesto JJ, Caru M (2002) Genetic diversity of Nostoc microsymbionts from Gunnera
tinctoria revealed by PCR-STRR fingerprinting. Microb Ecol 44:127–136
Gusev MV, Baulina OI, Gorelova OA, Lobakova ES, Korzhenevshaya TG (2002) Artificial
cyanobacterium-plant symbioses. In: Rai AN, Bergman B, Rasmussen U (eds) Cyanobacteria
in symbiosis. Kluwer, Dordrecht, pp 253–312
Herrero A, Muro-Pastor AM, Valladares A, Flores E (2004) Cellular differentiation and the NtcA
transcription factor in filamentous cyanobacteria. FEMS Microbiol Rev 28:469–487
Houle D, Gauthier SB, Paquet S, Planas D, Warren A (2006) Identification of two genera of N2fixing cyanobacteria growing on three feather moss species in boreal forests of Quebec,
Canada. Can J Bot 84:1025–1029
118
D.G. Adams and P.S. Duggan
Karthikeyan N, Prasanna R, Nain L, Kaushik BD (2007) Evaluating the potential of plant growth
promoting cyanobacteria as inoculants for wheat. Eur J Soil Biol 43:23–30
Karthikeyan N, Prasanna R, Sood A, Jaiswal P, Nayak S, Kaushik BD (2009) Physiological
characterization and electron microscopic investigation of cyanobacteria associated with
wheat rhizosphere. Folia Microbiol 54:43–51
Khamar HJ, Breathwaite EK, Prasse CE, Fraley ER, Secor CR, Chibane FL, Elhai J, Chiu W-L
(2010) Multiple roles of soluble sugars in the establishment of Gunnera-Nostoc endosymbiosis. Plant Physiol 154:1381–1389
Klint J, Ran L, Rasmussen U, Bergman B (2006) Identification of developmentally regulated
proteins in cyanobacterial hormogonia using a proteomic approach. Symbiosis 41:87–95
Kluge M (2002) A fungus eats a cyanobacterium: the story of the Geosiphon pyriformis
endocyanosis. Biol Environ Proc R Ir Acad 102B:11–14
Kluge M, Mollenhauer D, Wolf E, Shussler A (2002) The Nostoc-Geosiphon endocytobiosis. In:
Rai AN, Bergman B, Rasmussen U (eds) Cyanobacteria in symbiosis. Kluwer, Dordrecht,
pp 19–30
Knight CD, Adams DG (1996) A method for studying chemotaxis in nitrogen fixing cyanobacterium-plant symbioses. Physiol Mol Plant Pathol 49:73–77
Lechno-Yossef S, Nierzwicki-Bauer SA (2002) Azolla-Anabaena symbiosis. In: Rai AN, Bergman B,
Rasmussen U (eds) Cyanobacteria in symbiosis. Kluwer, Dordrecht, pp 153–178
Legaz M-E, Fontaniella B, Millanes A-M, Vicente C (2004) Secreted arginases from phylogenetically far-related species act as cross-recognition factors for two different algal cells. Eur J
Cell Biol 83:435–446
Lehr H, Galun M, Ott S, Jahns HM, Fleminger G (2000) Cephalodia of the lichen Peltigera
aphthosa (L.) Willd. Specific recognition of the compatible photobiont. Symbiosis 29:357–365
Liaimer A, Bergman B (2004) Phytohormones in cyanobacteria: occurrence and perspectives. In:
Tikhonovich I, Lugtenberg B, Provorov N (eds) Biology of plant-microbe interactions. International Society for Molecular Plant Microbe Interactions, St Paul, MN, pp 394–397
Liaimer A, Matveyev A, Bergman B (2001) Isolation of host plant induced cDNAs from Nostoc sp
strain PCC 9229 forming symbiosis with the angiosperm Gunnera spp. Symbiosis 31:293–307
Lindblad P (2009) Cyanobacteria in symbiosis with cycads. Microbiol Monogr 8:225–233
Lindblad P, Costa J-L (2002) The cyanobacterial-cycad symbiosis. Biol Environ Proc R Ir Acad
102B:31–33
Lindlbad P, Bergman B, Hofsten AV, H€allblom L, Nylund JE (1985) The cyanobacterium-Zamia
symbiosis: an ultrastructural study. New Phytol 101:707–716
Lobakova ES, Dol’nikova GA, Korzhenevshaya TG (2001a) Cyanobacterial-bacterial complexes
in plant syncyanoses. Microbiology 70:111–116
Lobakova ES, Shchelmanova AG, Korzhenevskaya TG, Gusev MV (2001b) Infection of plants
and plant tissue cultures with cyanobacteria-bacteria complexes. Microbiology 70:299–305
Lobakova ES, Orazova MK, Dobrovol’skaya TG (2003) Microbial complexes occurring on the
apogeotropic roots and in the rhizosphere of cycad plants. Microbiology 72:628–633
Mattick JS (2002) Type IV pili and twitching motility. Annu Rev Microbiol 56:289–314
Meeks JC (2003) Symbiotic interactions between Nostoc punctiforme, a multicellular cyanobacterium, and the hornwort Anthoceros punctatus. Symbiosis 35:55–71
Meeks JC (2006) Molecular mechanisms in the nitrogen-fixing Nostoc-bryophyte symbiosis. Prog
Molec Subcell Biol 41:165–196
Meeks JC (2009) Physiological adaptations in nitrogen-fixing Nostoc-plant symbiotic
associations. In: Pawlowski K (ed) Prokaryotic symbionts in plants, vol 8, Microbiol
monographs. Springer, Berlin, pp 181–205
Meeks JC, Elhai J (2002) Regulation of cellular differentiation in filamentous cyanobacteria in
free-living and plant-associated symbiotic growth states. Microbiol Mol Biol Rev 66:94–121
Meeks JC, Elhai J, Thiel T, Potts M, Larimer F, Lamerdin J, Predki P, Atlas R (2001) An overview
of the genome of Nostoc punctiforme, a multicellular, symbiotic cyanobacterium. Photosynth
Res 70:85–106
Signalling in Cyanobacteria–Plant Symbioses
119
Meeks JC, Campbell EL, Summers ML, Wong FC (2002) Cellular differentiation in the cyanobacterium Nostoc punctiforme. Arch Microbiol 178:395–403
Muro-Pastor MI, Reyes JC, Florencio FJ (2001) Cyanobacteria perceive nitrogen status by sensing
intracellular 2-oxoglutarate levels. J Biol Chem 276:38320–38328
Muro-Pastor AM, Reyes JC, Florencio FJ (2005) Ammonium assimilation in cyanobacteria.
Photosynth Res 83:135–150
Nilsson MC, Wardle DA (2005) Understory vegetation as a forest ecosystem driver: evidence from
the northern Swedish boreal forest. Eur J Phycol 3:421–428
Nilsson M, Bergman B, Rasmussen U (2000) Cyanobacterial diversity in geographically related
and distant host plants of the genus Gunnera. Arch Microbiol 173:97–102
Nilsson M, Bhattacharya J, Rai AN, Bergman B (2002) Colonization of roots of rice (Oryza sativa)
by symbiotic Nostoc strains. New Phytol 156:517–525
Nilsson M, Rasmussen U, Bergman B (2005) Competition among symbiotic cyanobacterial
Nostoc strains forming artificial associations with rice (Oryza sativa). FEMS Microbiol Lett
245:139–144
Nilsson M, Rasmussen U, Bergman B (2006) Cyanobacterial chemotaxis to extracts of host and
nonhost plants. FEMS Microbiol Ecol 55:382–390
Nudleman E, Kaiser D (2004) Pulling together with type IV pili. J Mol Microbiol Biotechnol
7:52–62
Osborne B, Sprent JI (2002) Ecology of the Nostoc-Gunnera symbiosis. In: Rai AN, Bergman B,
Rasmussen U (eds) Cyanobacteria in symbiosis. Kluwer, Dordrecht, pp 233–251
Pabby A, Prasanna BM, Nayak S, Singh AK (2003) Physiological characterization of the cultured
and freshly isolated endosymbionts from different species of Azolla. Mar Biotechnol 41:73–79
Pabby A, Prasanna BM, Singh AK (2004a) Biological significance of Azolla and its utilization in
agriculture. Proc Indian Natl Sci Acad B 3:299–333
Pabby A, Prasanna R, Singh PK (2004b) Morphological characterization of cultured and freshly
separated cyanobionts (Nostocales, Cyanophyta) from Azolla species. Acta Bot Hung 46:211–223
Papaefthimiou D, Hrouzek P, Mugnai MA, Lukesova A, Turicchia S, Rasmussen U, Ventura S
(2008a) Differential patterns of evolution and distribution of the symbiotic behaviour in
nostocacean cyanobacteria. Int J Syst Evol Microbiol 58:553–564
Papaefthimiou D, Van Hove C, Lejeune A, Rasmussen U, Wilmotte A (2008b) Diversity and host
specificity of Azolla cyanobionts. J Phycol 44:60–70
Paulsrud P, Lindblad P (2002) Fasciclin domain proteins are present in Nostoc symbionts of
lichens. Appl Environ Microbiol 68:2036–2039
Rai AN, Soderback E, Bergman B (2000) Cyanobacterium-plant symbioses. New Phytol
147:449–481
Ran LA, Larsson J, Vigil-Stenman T, Nylander JAA, Ininbergs K, Zheng WW, Lapidus A, Lowry
S, Haselkorn R, Bergman B (2010) Genome erosion in a nitrogen-fixing vertically transmitted
endosymbiotic multicellular cyanobacterium. PLoS One 5:e11486
Rasmussen U, Nilsson M (2002) Cyanobacterial diversity and specificity in plant symbioses. In:
Rai AN, Bergman B, Rasmussen U (eds) Cyanobacteria in symbiosis. Kluwer, Dordrecht,
pp 313–328
Rasmussen U, Svenning MM (2001) Characterization by genotypic methods of symbiotic Nostoc
strains isolated from five species of Gunnera. Arch Microbiol 176:204–210
Rasmussen U, Johansson C, Renglin A, Petersson C, Bergman B (1996) A molecular characterization of the Gunnera-Nostoc symbiosis: comparison with Rhizobium- and Agrobacteriumplant interactions. New Phytol 133:391–398
Renzaglia KS, Duff RJ, Nickrent DL, Garbary D (2000) Vegetative and reproductive innovations
of early land plants: implications for a unified phylogeny. Philos Trans R Soc Lond B Biol Sci
355:769–793
Renzaglia KS, Schuette S, Duff RJ, Ligrone R, Shaw AJ, Mishler BD, Duckett JG (2007)
Bryophyte phylogeny: advancing the molecular and morphological frontiers. Bryologist
110:179–213
120
D.G. Adams and P.S. Duggan
Rikkinen J (2002) Cyanolichens: an evolutionary overview. In: Rai AN, Bergman B, Rasmussen U
(eds) Cyanobacteria in symbiosis. Kluwer, Dordrecht, pp 31–72
Rikkinen J, Virtanen V (2008) Genetic diversity in cyanobacterial symbionts of thalloid
bryophytes. J Exp Bot 59:1013–1021
Sacristan M, Millanes A-M, Legaz M-E, Vicente C (2006) A lichen lectin specifically binds to the
alpha-1,4-polygalactoside moiety of urease located in the cell wall of homologous algae. Plant
Signal Behav 1:23–27
Seifert GJ, Roberts K (2007) The biology of arabinogalactan proteins. Annu Rev Plant Biol
58:137–161
Sergeeva E, Liaimer A, Bergman B (2002) Evidence for production of the phytohormone indole-3acetic acid by cyanobacteria. Planta 215:229–238
Showalter AM (2001) Arabinogalactan proteins: structure, expression and function. Cell Mol Life
Sci 58:1399–1417
Solheim B, Zielke M (2002) Associations between cyanobacteria and mosses. In: Rai AN,
Bergman B, Rasmussen U (eds) Cyanobacteria in symbiosis. Kluwer, Dordrecht, pp 137–152
Solheim B, Wiggen H, Roberg S, Spaink HP (2004) Associations between arctic cyanobacteria
and mosses. Symbiosis 37:169–187
Sood A, Prasanna R, Prasanna BM, Singh PK (2008a) Genetic diversity among and within cultured
cyanobionts of diverse species of Azolla. Folia Microbiol 53:35–43
Sood A, Prasanna R, Singh PK (2008b) Fingerprinting of freshly separated and cultured
cyanobionts from different Azolla species using morphological and molecular markers.
Aquat Bot 88:142–147
Svenning MM, Eriksson T, Rasmussen U (2005) Phylogeny of symbiotic cyanobacteria within the
genus Nostoc based on 16S rDNA sequence analyses. Arch Microbiol 183:19–26
Thajuddin N, Muralitharan G, Sundaramoorthy M, Ramamoorthy R, Ramachandran S, Akbarsha
MA, Gunasekaran M (2010) Morphological and genetic diversity of symbiotic cyanobacteria
from cycads. J Basic Microbiol 50:254–265
Uheda E, Silvester WB (2001) The role of papillae during the infection process in the GunneraNostoc symbiosis. Plant Cell Physiol 42:780–783
Ungerer JL, Pratte BS, Thiel T (2008) Regulation of fructose transport and its effect on fructose
toxicity in Anabaena spp. J Bacteriol 190:8115–8125
Usher KM (2008) The ecology and phylogeny of cyanobacterial symbionts in sponges. Mar Ecol
Evol Perspect 29:178–192
Vaishampayan A, Sinha RP, Hader D-P, Dey T, Gupta AK, Bhan U, Rao AL (2001)
Cyanobacterial biofertilizers in rice agriculture. Bot Rev 67:453–516
van Hove C, Lejeune A (2002a) The Azolla-Anabaena symbiosis. Biol Environ Proc R Ir Acad
102B:23–26
van Hove C, Lejeune A (2002b) Applied aspects of Azolla-Anabaena symbiosis. In: Rai AN,
Bergman B, Rasmussen U (eds) Cyanobacteria in symbiosis. Kluwer, Dordrecht, pp 179–193
Vazquez-Bermudez MF, Herrero A, Flores E (2002) 2-oxoglutarate increases the binding affinity
of the NtcA (nitrogen control) transcription factor for the Synechococcus glnA promoter. FEBS
Lett 512:71–74
Vessey JK, Pawlowski K, Bergman B (2005) Root-based N2-fixing symbioses: legumes,
actinorhizal plants, Parasponia sp. and cycads. Plant Soil 274:51–78
Villarreal JC, Renzaglia KS (2006) Structure and development of Nostoc strands in
Leiosporoceros dussii (Anthocerotophyta): a novel symbiosis in land plants. Am J Bot 93:
693–705
Vivas M, Sacristan M, Legaz M-E, Vicente C (2010) The cell recognition model in chlorolichens
involving a fungal lectin binding to an algal ligand can be extended to cyanolichens. Plant Biol
12:615–621
Wang CM, Ekman M, Bergman B (2004) Expression of cyanobacterial genes involved in
heterocyst differentiation and dinitrogen fixation along a plant symbiosis development profile.
Mol Plant Microbe Interact 17:436–443
Signalling in Cyanobacteria–Plant Symbioses
121
Watts SD (2000) Signalling in the Nostoc-plant symbiosis. PhD thesis, University of Leeds, UK,
pp 185
West NJ, Adams DG (1997) Phenotypic and genotypic comparison of symbiotic and free-living
cyanobacteria from a single field site. Appl Environ Microbiol 63:4479–4484
Wong FCY, Meeks JC (2001) The hetF gene product is essential to heterocyst differentiation and
affects HetR function in the cyanobacterium Nostoc punctiforme. J Bacteriol 183:2654–2661
Wong FCY, Meeks JC (2002) Establishment of a functional symbiosis between the cyanobacterium Nostoc punctiforme and the bryophyte Anthoceros punctatus requires genes involved in
nitrogen control and initiation of heterocyst differentiation. Microbiology 148:315–323
Wouters J, Janson S, Bergman B (2000) The effect of exogenous carbohydrates on nitrogen
fixation and hetR expression in Nostoc PCC 9229 forming symbiosis with Gunnera. Symbiosis
28:63–76
Zhang C-C, Laurent S, Sakr S, Peng L, Bedu S (2006) Heterocyst differentiation and pattern
formation in cyanobacteria: a chorus of signals. Mol Microbiol 59:367–375
Zheng WW, Song TY, Bao XD, Bergman B, Rasmussen U (2002) High cyanobacterial diversity in
coralloid roots of cycads revealed by PCR fingerprinting. FEMS Microbiol Ecol 40:215–222
Zheng WW, Ran L, Bergman B (2009a) Structural characteristics of the cyanobacterium-Azolla
symbiosis. In: Pawlowski K (ed) Prokaryotic symbionts in plants, vol 8, Microbiol
monographs. Springer, Heidelberg, pp 235–263
Zheng WW, Bergman B, Chen B, Zheng SP, Xiang G, Rasmussen U (2009b) Cellular responses in
the cyanobacterial symbiont during its vertical transfer between plant generations in the Azolla
microphylla symbiosis. New Phytol 181:53–61
Zielke M, Ekker AS, Olsen RA, Spjelkavik S, Solheim B (2002) The influence of abiotic factors on
biological nitrogen fixation in different types of vegetation in the High Arctic, Svalbard. Arct
Antarct Alpine Res 34:293–299
Zielke M, Solheim B, Spjelkavik S, Olsen RA (2005) Nitrogen fixation in the high arctic: role of
vegetation and environmental conditions. Arct Antarct Alpine Res 37:372–378
Signalling in Ectomycorrhizal Symbiosis
Judith Felten, Francis Martin, and Valérie Legué
Abstract The mechanism by which tree roots and soil fungi interact and form their
common, symbiotic organ, the ectomycorrhiza (ECM), involves numerous steps.
During this ontogenic process, the developmental programs of both partners are
modified in order to enable symbiosis establishment. Both roots and fungus release
an array of various metabolites (morphogens and signalling molecules) that establish a molecular cross-talk between symbionts. In contrast to some other
plant–microbe interactions, such as rhizobia or arbuscular mycorrhiza symbiosis,
the characterization of these signalling molecules and their impact on developmental pathways is poorly known. Recent studies have provided new insights into
specific phases and signalling pathways of ECM development on a molecular
level and have thereby started to fill the gaps in our understanding of root–fungus
communication. Based on this knowledge and recent data from ECM interaction, we
will identify possible crosstalk between ECM signalling and root development.
1 Structure and Development of Ectomycorrhizae
While only around 3% of seed-bearing plants establish an ectomycorrhizal (ECM)
symbiosis with fungi, ECMs are the most common form of symbiosis for the
majority of boreal and temperate forest trees. The fungal community involved in
J. Felten
UMR 1136 Interactions Arbres Micro-organismes, Centre INRA de Nancy,
54280 Champenoux, France
Department of Forest Genetics and Plant Physiology, Umeå Plant Science Centre, Swedish
University of Agricultural Sciences, 90 187 Umeå, Sweden
F. Martin • V. Legué (*)
UMR 1136 Interactions Arbres Micro-organismes, Centre INRA de Nancy,
54280 Champenoux, France
e-mail: [email protected]; [email protected]
S. Perotto and F. Baluska (eds.), Signaling and Communication in Plant Symbiosis,
Signaling and Communication in Plants 11, DOI 10.1007/978-3-642-20966-6_6,
# Springer-Verlag Berlin Heidelberg 2012
123
124
J. Felten et al.
ECM formation concerns a great diversity of species among Basidiomycota and
Ascomycotina fungi (Smith and Read 2008). Plants species forming ECM belong to
Pinophyta and Angiosperms in temporate, boreal, and partially tropical forest
(Smith and Read 2008).
The crucial impact of ECM fungi on tree nutrient uptake, including uptake of
major nutrients such as nitrogen (N), phosphorus (P), and magnesium (Mg), is well
established (reviews Chalot et al 2006; Martin and Nehls 2009). In parallel, the
change in tree carbon (C) allocation caused by ECM fungi is considerable. These
metabolism activities confer a major role of ECM in ecological and biogeochemical
processes in forests (Taylor 2002; Buée et al. 2009).
The functioning structure defined as an ECM is characterized by a basic pattern
including the presence of a typical mantle of fungal hyphae around the root, and a
labyrinthine inward growth of hyphae between epidermal and (in some species)
cortical cells, called the Hartig net (Blasius et al. 1986; Fig. 1a), while the intracellular penetration is scarce. An outwardly growing network of hyphal elements, the
extramatrical mycelium, is seasonal present. This well-conserved feature drives the
nutrient flow between the two partners (Martin 2007). Extramatrical hyphae gather
nutrients (mostly N and P) from the soil and transport them to mantle hyphae, where
they may be stored. From there, nutrients are transported to Hartig net hyphae and
delivered to and taken up by root epidermis (and cortex) cells. Vice versa,
Fig. 1 Anatomy and structure of ectomycorrhizae. (a) Populus tremula x Populus alba/Laccaria
bicolor root cross section. Fluorescent double staining showing peripheric root cells (red cell wall)
and fungal hyphae (green cell wall). Mantle (M), Hartig Net (HN), extramatrical hyphae (EH),
root epidermis cells (EC), root cortex cells (CC). Bar 10 mm. (b) Ectomycorrhizal Poplar/Laccaria
bicolor short roots. (c) Poplar root apex colonized by Laccaria bicolor. The mantle (M) is present
in the root apex, but the Hartig net is developed behind the root meristem. The ECM is
accompanied with a reduced root cap (RC) and a small apical meristem zone. Bar 20 mm.
(d) Interface (I) between root cells (RC) and fungal cells (FC) inside the Hartig net. Courtesy
J. Gérard (Nancy University)
Signalling in Ectomycorrhizal Symbiosis
125
photoassimilates (sugars) are released by plant root cells and taken up by the fungus
before (partially) being transported back into extramatrical hyphae that use them for
further growth and fruiting body development. Together, the different structures
and nutrient flow stimulate the growth and health of the plant on several levels: (1)
thanks to the wide network of extramatrical hyphae, it benefits from nutrient
resources from a greater soil volume than it could exploit by its roots alone
(Rousseau et al. 1994), (2) the secreted enzyme activities, which increase mineral
solubilization in the soil, can enhance nutrient absorption by plant roots
(Landeweert et al. 2001), and (3) the enclosure of the root by the fungus protects
plant roots against pathogens but also against inorganic agents. In specific cases, the
ability of fungi to absorb and store heavy metals (Blaudez et al. 2000) can even
open an ecological niche for the plant that it would not be able to grow in without
being associated with ECM fungi.
Like in the formation of endomycorrhizae, ECM development involves a series
of complex events. To elucidate such events, most research has employed
simplified in vitro systems that are conducted to separate sequential phases
(Horan et al. 1988; Martin and Tagu 1999; Fig. 2). In the first precontact phase,
commonly called early phase, the fungus is attracted towards the root from a soil
propagule by increasing hyphal branching and directional growth or from spores by
inducing their germination. Fungal hyphae progressively colonize and adhere to the
root epidermis (Fig. 2c, d). The docking process that consists of hyphae attachment
to root epidermal cells starts with the formation of an “adhesion pad” through
aggregation of hyphae (Jacobs et al. 1989). It has been proposed that the initial
interaction of hyphae and roots during colonization occurs close to the root tip and
that subsequently hyphae invade the root from root cap cells in- and upwards to the
epidermis (Horan et al. 1988). Depending on the species, either a mantle of multiple
layers of hyphae that multiply and differentiate (Horan et al. 1988) forms first
around the root from which the Hartig net develops inward, or occasionally the
Hartig net can establish first and the mantle forms afterwards (Nylund and Unestam
1982).
Once colonization of a certain root part is accomplished and the mantle and
Hartig net are well developed, the nutrient exchanges take place between both
partners in the Hartig net, leading to the functional mycorrhiza, called late phase
(Fig. 2e). Furthermore, an equilibrium is established in terms of colonization:
depending on plant and fungal species, the Hartig net attains a certain depth into
the cell layers of the root (from the epidermis cells to cortex cells) and once
colonization is accomplished the fungus does not penetrate any further. The equilibrium (biotrophic) phase, where efficient nutrient exchange occurs, may be
maintained for a certain period of time, on the species involved (Smith and Read
2008), until the mycorrhiza ages and undergoes senescence (Dexheimer et al.
1986). It has been shown that ageing mycorrhizas may lose their fungal mantle,
even if the Hartig net is maintained, and that their contribution to plant–fungus
nutrient exchange diminishes (Al-Abras et al. 1988). The fungus can return back to
a saprophytic lifestyle and invade senescent root cells (necrotrophic phase).
126
J. Felten et al.
Fig. 2 Scheme of ECM development. ECM development can be separated into three main phases.
(1) Signal exchange prior to physical contact, (2) colonization, and (3) nutrient exchange in the
functional ECM. The precontact phase is characterized by a plant-to-fungus signalling to induce
hyphae branching and attract the fungus (a) and to a fungus-to-plant signalling, which stimulates
lateral root development of the plant (b). The colonization phase may be separated into a first
recognition and adhesion of hyphae on the root (c) and of penetration of fungal hyphae in between
epidermal (and cortex) cells of the plant (d), represented as a transverse root section. The
functional ectomycorrhiza is characterized by active nutrient exchange (ammonium (N) and
phosphate (P) from the fungus to the plant and sugars (C) from the plant to the fungus).
Extramatrical hyphae explore nutrients in the soil and transport them to the mantle and Hartig
Net where specific transporters insure the exchange
Signalling in Ectomycorrhizal Symbiosis
127
2 Reprogramming of Root Development
One of the observations when ECM were first described was the presence of
numerous short and swollen lateral roots all over the root system (Fig. 1b). The
presence of this type of root only during symbiosis illustrates the impact of the
fungus on root system development.
The increase in lateral root formation is common for all studied ECM systems.
Interestingly, these changes have been observed in response to endomycorrhizae
fungi and also in response to bacteria (Olah et al. 2005), suggesting some common
traits in the response of the plant host. Moreover, a stimulation of lateral root
formation occurs during a change in nutrient availability, including N and P,
suggesting a link between the mineral absorption by the fungus during the establishment of ECM and their effects on root architecture.
Recently, it has been observed in an in vitro system that the first visible modification in the induction of lateral roots, which emerge in response to fungal signals,
from the parental roots starts in the very early phase of plant–fungus contact, even
prior to physical contact between partners (Splivallo et al. 2009; Felten et al. 2009).
These observations suggest the importance of a diffusible and/or a volatile signals
in this change.
In the more advanced phases of ECM development, another striking change in
root development can be observed, i.e., the formation of short roots. The detailed
anatomical observations in Eucalyptus globulus in contact with Pisolithus clearly
reveal a reduced root cap and a small apical meristem zone (Massicote et al. 1987).
The differentiation of root tissues including endodermis, stele, and cortex starts
closer to the apex. Our observations in poplar–Laccaria bicolor follow a broadly
similar pattern to that of Eucalyptus (Fig. 1c). In conifer species, dichotomy of the
root apical meristem can be observed, which results in short roots that branch just
below their tip (Dexheimer and Pargney 1991; Laajanen et al. 2007).
More recently, it has been shown that root growth is guaranteed by the balance
between stem cell proliferation and differentiation leading to a specific pattern of
root organization. An auxin gradient, dictated mainly by auxin transport and the
combinatorial action of some transcription factors, maintains the cell stem niche
(Terpstra and Heidstra 2009). Interestingly, the auxin gradient is modified during
the first stage of ECM formation (Felten et al. 2009). However, the potential
function of transcription factors in the context of ECM is not established and fungal
signalling molecules acting on the root apical meristem and columella initials are
currently under investigation.
Another striking change in root development is the decay of root hairs in ECM
(Ditengou et al. 2000). The absence of root hairs is thought to be due to the physical
constraints of the fungal mantle around the root. Root hairs in plants are suggested
to fulfill functions in nutrient (N, P) absorption (Bates and Lynch 2000). During
symbiosis extramatrical hyphae efficiently take over these functions, so that root
hairs are likely to become unimportant for proper ECM root nutrient provisioning.
128
J. Felten et al.
However, it remains to be elucidated how the root developmental program is altered
to halt development of root hairs.
3 Remodelling of Root Cell Walls
Cell walls are important features of plant cells that perform a number of essential
functions, including providing shape to the many different cell types. Forming the
interface between adjacent cells, plant cell walls often play important roles in
intercellular communication. Because of their surface location, plant cell walls
also play an important role in plant–microbe interactions.
In the context of ECM, hyphae aggregation and formation of the adhesion pad at
the root surface involves recognition between both partners and synthesis of
different cell surface structures. The formation of the new interface between the
two partners plays an important role in the maintenance and the functioning of the
ECM. Ultrastructural studies (Dexheimer and Pargney 1991) reveal that the interface is not formed by the superposition of the fungal wall and the cortical cell walls
of the host plant. The parietal structure of the partners is modified to some degree,
with, as a result, the production of a polysaccharide-rich cement involved in the
cohesion of both partners and reinforcing the anchorage of the fungus to the plant
cell surface (Fig. 1d). The presence of chitin has not been detected in this interface.
This extensive modification of root cell walls is correlated with the induction of
genes encoding cell wall synthesis, loosening and degrading enzymes that may
facilitate the entrance of fungi between root cells (Duplessis et al. 2005). The same
mycelium can colonize either the epidermal layer only or the cortical layer as well,
depending on the host plant. This demonstrates that the host plant controls the
fungal growth habit, but the molecular mechanisms are unknown.
Certain fungal cell wall polypeptides are specifically expressed in Pisolithus
before or during early phases of Eucalyptus root colonization and are downregulated later in the mature mycorrhiza (Duplessis et al. 2005; Hilbert et al.
1991; Hilbert and Martin 1988; Martin and Tagu 1995). These are termed “Symbiosis Related Acidic Polypeptides” (SRAPs). Another group of cell wall-related
proteins that were overexpressed specifically at the same time points in
P. microcarpus were hydrophobins. These molecules secreted by the fungus are
considered to facilitate the intercellular penetration of hyphae (Nylund 1980) and
consequently to have a considerable role in the construction of this new interface
and in the communication between the two partners. Hydrophobins are small
hydrophobic polypeptides with a conserved distribution of eight cystein residues.
Only one class (class I) is found in Basidiomycetes (Wessels 1997). The immunolocalization of one of these hydrophobins, named HYDPt-1, from Pisolithus
tinctorius (Tagu et al. 2001) reveals its presence in the surface of the hyphae in
the mantle and in the Hartig net. The role of hydrophobins in ECM is not well
known. It has been proposed that these proteins facilitate the communication
between the two partners in participating in the construction of the surface.
Signalling in Ectomycorrhizal Symbiosis
129
In parallel, the establishment of ECM is accompanied by profound modifications
in the host cortical cells walls. The orientation of cell growth is radial, suggesting a
transformation of the parietal morphogenesis. However, no studies describe in
detail the mechanisms involved.
These amazing processes occurring in the host root development implicate a
complex signalling network between the two partners.
4 The ECM Fungus Perceives Plant Signals
It has been suggested that root exudates contain compounds that are recognized by
fungi and that attract them (Horan and Chilvers 1990). In Eucalyptus, roots release
the flavonol rutin that has been identified to stimulate growth of the fungus P.
tinctorius at only a picomolar in concentration (Lagrange et al. 2001; Martin et al.
2001). It was furthermore shown that the cytokinin zeatin is able to induce
branching in ECM hyphae (Martin et al. 2001; Gogala 1991). These factors might
be the first communication between a plant and its symbiotic ECM fungus in the
early phase (Fig. 2). Their attribution to the early phase can be made because it was
shown that when a barrier is present between roots and fungi, compatible fungi will
recognize the root and grow towards it (Horan and Chilvers 1990), suggesting that
molecules present in root exudates are sufficient for this step. The fact that a
flavonol is included in these “branching factors” is an interesting finding, as
rhizobia bacteria are also attracted by flavonoids secreted by the plant (Aguilar
et al. 1988). This indicates that similarities may occur between different symbioses
concerning attraction of the microbial symbiont by the plant partner. Interestingly,
the perception of plant exudates not only triggers development of hyphae but also
accumulation of metabolites, such as hypaphorine, which may be involved in
fungus to plant signalling and will be discussed below (Beguiristain and Lapeyrie
1997). Abietic acid extracted from Pinus roots was able to induce spore germination
at a very low concentration (10 7 M) and this effect seems to be specific to the
genus Suillus (Fries et al. 1987). Horan and Chilvers (1990) demonstrated that the
presence of root-diffusible molecules is able to chemioattract ECM mycelia.
In the case of AM interactions, studies have revealed the role of strigolactone as
a signalling compound to the hyphae aiding in branching and establishment of the
mycorrhizae (Gomez-Roldan et al. 2008). Chemical signals released by symbiotic
bacteria and by AM fungi are characterized; however, similar molecules have not
been identified in ECM.
5 Hormone Signalling During ECM Development
Phytohormones (plant-growth substances) are metabolites that exist in extremely
low quantities in plants, or also in microorganisms, and influence in a dosedependent manner the development of all plant organs. In addition to the five
130
J. Felten et al.
“classical phytohormones” auxin, ethylene, gibberellins, abscisic acid, and
cytokinins (Kende and Zeevaart 1997), today, further molecules such as salicylic
acid, brassinosteroids (Grove et al. 1979), and jasmonic acid (Creelman and Mullet
1997) have been assigned as phytohormones and further growth-modifying
substances [e.g. strigolactones (Gomez-Roldan et al. 2008; Umehara et al. 2008)]
are discussed as having phytohormone character.
In roots, different phytohormones are involved in lateral root development
(Fukaki and Tasaka 2009), root meristem maintenance (Benkova and Hejatko
2009), root hair development, and elongation (Rahman et al. 2002; Pitts et al.
1998; Jones et al. 2009) and formation of the root cap (Wang et al. 2005). Auxin
is a central element in root development, but, via crosstalk with auxin, other
phytohormones also influence root development (Fukaki and Tasaka 2009;
Benkova and Hejatko 2009). The fact that during ECM symbiosis root development
is altered on all levels suggests that fungal signalling molecules can interact with or
alter the root auxin levels that control these patterns. Because of their versatile
nature and their production by plants and fungi, phytohormones are interesting
candidates to mediate plant–fungus signalling during ECM development.
5.1
Volatile Phytohormones as Precontact Signals
Different studies have suggested that the first fungal signal perceived by the plant
does not require any physical contact with the fungus (Splivallo et al. 2009; Felten
et al. 2009, 2010). This conclusion has been based on studies that have
demonstrated that LR formation in mycorrhizal or nonmycorrhizal plants is
stimulated even when ECM truffle fungi or L. bicolor are spaced at a certain
distance in a Petri dish, or are separated by a semipermeable barrier, from the
plant. More strikingly, LR stimulation will even occur when only the exchange of
volatiles and not soluble molecules between both partners is allowed (Felten et al.
2010). Therefore, the hypothesis was established that volatile molecules released by
the fungus may be the first fungal signal(s) perceived by the plant. ECM fungi are
able to produce volatiles such as the phytohormone ethylene (Rupp et al. 1989;
Graham and Linderman 1980; Splivallo et al. 2009). Others have demonstrated that
the fungus Fusarium oxysporum (a nonectomycorrhizal fungus) releases different
jasmonates (Miersch et al. 1999). As diffusible molecules such as ethylene and
jasmonates are known for their stimulatory effect on LR development (Regvar et al.
1997; Sun et al. 2009; Ivanchenko et al. 2008), they may be some of the key actors
in fungal-induced LR stimulation during the precolonization stage. Splivallo et al.
(2009) considered ethylene released by ECM truffle fungi as a stimulator of LR
development in Arabidopsis as LR stimulation by these fungi was decreased in the
ethylene insensitive Arabidopsis line ein2. However, in feeding experiments ethylene mimicked fungal LR induction only when applied together with exogenous
auxin. Moreover certain ECM fungi, such as P. tinctorius, do not produce ethylene
but nevertheless stimulate LR development (Rupp et al. 1989). This suggests that
Signalling in Ectomycorrhizal Symbiosis
131
ethylene may be part of the LR stimulating signals exchanged between fungus and
plant during the early phase of interaction, but that it is not the only one. Interestingly, the exogenous application of methyljasmonate accelerated the first mycorrhizal contact of spruce roots and L. laccata (Regvar et al. 1997), which also
suggests a possible role for methyljasmonate during the early phases of ECM
establishment. It will be interesting to analyze whether jasmonates are produced
by ECM fungi and to further study their effect on ECM development.
It is worth noting that experiments on ethylene levels during colonization have
not distinguished between ethylene derived from the plant or from the fungus. The
finding of Rupp et al. (1989) that there was an increase in ethylene levels released
by roots colonized either with L. bicolor (produces ethylene) or with P. tinctorius
(does not produce ethylene) suggests that the plant partner produces ethylene upon
colonization. The same could be valid for jasmonates. Jasmonate and ethylene are
usually associated with defense responses against necrotrophic pathogens or herbivore insects (Bari and Jones 2009; Plett 2010), but act as well in biotrophic
(mutualistic) associations (Gutjahr and Paszkowski 2009). Consequently, they
may be involved in the early defense response during root–ECM fungus contact.
5.2
The Role of Auxin During Early Contact Phase
Auxin, which is one of the major regulators of root development, has been
suggested to be at the crosspoint of fungus–plant signalling and the modification
of root development during ECM establishment (Felten et al. 2009; Splivallo et al.
2009; Laajanen et al. 2007; Raudaskoski and Salo 2008).
Slankis (1950) was one of the first to suggest a function for auxin released by
ECM fungi in the development of ECM root tips. Since then, different research
groups have addressed auxin production by ECM fungi in axenic cultures.
Techniques ranging from thin-layer chromatography experiments (Ho 1987a, b)
to quantification with ELISA techniques or IAA antibodies (Karabaghli-Degron
et al. 1998; Rincon et al. 2003) and GC–MS analysis (Splivallo et al. 2009) have all
found a very low production of fungal auxin in the nanomolar range (10–300 nM)
for different fungi (Ho 1987a, b; Rincon et al. 2001, 2003; Reddy et al. 2006).
While results from these studies indicate a large difference in the ability of
individual fungal strains to produce IAA (Ek et al 1983; Rudawska and
Kieliszewska-Rokicka 1997), even the highest amount of auxin released by certain
ECM fungi appears too low to be responsible for early plant responses such as
LR stimulation. This is confirmed by the results of Karabaghli-Degron et al. (1998),
who demonstrated that LR induction in Norway spruce required 100–500 mM
exogenously applied IAA and that below this concentration no effect was observed.
However challenging roots with L. bicolor, which secretes only about 10 nM IAA,
resulted in a visible LR stimulation. Splivallo et al. (2009) came to similar
conclusions. They revealed an induction of the auxin response in roots during
contact with truffle fungi, as visualized by Arabidopsis thaliana pDR5: GFP
132
J. Felten et al.
lines. However, the fungi only released about 100 nM IAA into the medium, which
when exogenously applied did not reproduce the effect of the fungus. Recently, we
were able to show that when auxin and L. bicolor are simultaneously applied to A.
thaliana, a cumulative effect is observed concerning LR stimulation, suggesting
that fungal signals and auxin can work in synergy to induce LRs (Felten et al. 2010).
Taken together these results highlight that fungal auxin in its very low quantity is
unlikely to be the trigger of the early plant–fungus signalling. But what about
internal root auxin?
We addressed this question in a study where we analyzed whether functional
polar auxin transport inside poplar and A. thaliana roots was necessary for the
fungus to stimulate LR development (Felten et al. 2009). Polar auxin transport
occurs due to specific auxin efflux carriers (PIN proteins) situated at distinct sides of
the plasma membrane of a cell that mediate the release of auxin from the cell. This
directed transport permits auxin to accumulate at specific sites in the root, where it
triggers, for example, lateral root primordium initiation or meristem maintenance
(Fukaki and Tasaka 2009). Our results showed that polar auxin transport is required
for the fungus to stimulate LR formation in poplar or Arabidopsis plants. Interestingly, one specific member of the PIN multigene family, PtPIN9 (orthologue of
AtPIN2) was identified as necessary for fungus-induced LR stimulation. Interestingly, this member of the PIN family in Arabidopsis is responsive to jasmonates
(Sun et al. 2009). PIN2 induction by the presence of the fungus during the
precontact phase is therefore another step towards implication of volatile fungal
molecules during the early fungus–plant signalling.
Moreover via their crosstalk with auxin, jasmonates and ethylene are also able to
directly influence auxin biosynthesis in plant tissues (Fukaki and Tasaka 2009;
Benkova and Hejatko 2009; Stepanova et al. 2007). The observed enhanced auxin
response in roots during plant–fungus contact (Splivallo et al. 2009) may therefore
be a secondary effect of directed auxin transport or auxin synthesis, not an accumulation of fungal auxin inside plant roots. In conclusion, fungal auxin is likely to play
a minor role during the early signalling phase whereas other versatile signals, such
as phytohormones ethylene and jasmonates, are likely to indirectly enhance auxin
responses inside the root (Fig. 3).
5.3
Auxin as a Possible Signal During Cell Loosening
and Hartig Net Development
Low amounts of auxin released by the fungus may be of major importance for root
colonization once both partners come into close proximity and released auxin can
accumulate inside plant tissues. As part of the “Acid-Growth Theory,” auxin, as a
weak acid, has been reported to have the ability to loosen the cell wall and to permit
growth of cells (Rayle and Cleland 1992). Furthermore, Mensen et al. (1998)
proposed that fungal auxin could reduce the peroxidase-catalyzed linkage of the
Signalling in Ectomycorrhizal Symbiosis
133
Fig. 3 Auxin homeostasis during ECM development
cell wall constituents, allowing hyphae to penetrate between the cortical cells to
form the Hartig net. Results from Tranvan et al. (2000) illustrated nicely how IAA
overproduction by the fungus Hebeloma cylindrosporum resulted in a quicker and
deeper colonization of the root.
Reddy et al. (2006) as well as Charvet-Candela et al. (2002) had reported the
differential expression of auxin-inducible genes PpGH3-16 and PpIAA88 in Pinus
pinaster roots during colonization by H. cylindrosporum or Rhizopogon roseolus.
Interestingly, their studies demonstrated no difference in transcript amount between
roots challenged with wild type or the IAA-overproducing strain but a faster
transcript change when the auxin overproducer was used. This indicates that the
134
J. Felten et al.
accumulation of fungal auxin inside plant tissues triggers the rapidity and the final
outcome of colonization.
In our own study (Felten et al. 2009; Fig. 3) we detected in poplar roots during
colonization an accumulation of transcripts related to enzymes (IAA-amido-synthetases
GH3) that are able to conjugate auxin to amino acids and thereby mediate
dynamically auxin homeostasis (Staswick et al. 2005) and of auxin influx carriers,
which import auxin into the cell. It may hence be possible that the plant actively
removes fungal auxin from the apoplast and re-adjusts auxin levels. One reason for
this could be to divert the free auxin into inactive conjugates or to degrade it.
Indeed, Wallender et al. (1992) observed a decrease in auxin levels in mature ECM
compared to noncolonized plants. This may be a result of the balance of auxin
production by the fungus and auxin degradation by the plant in mature ECM where
auxin levels may be drastically reduced to restrict further penetration deeper into
plant tissues. Thus, auxin-homeostasis adjustment could be part of the mechanism
that regulates the equilibrium of Hartig net colonization (Figs. 3 and 4).
Lastly, Karabaghli-Degron et al. (1998) and Rincon et al. (2001) addressed
whether inhibition of polar auxin transport (acting on the plant and apparently not
on auxin secretion from the fungus) impacted root colonization. They discovered
that the polar auxin transport inhibitor TIBA completely inhibited Eucalyptus root
colonization by Pisolithus and the inhibitor NPA caused an irregular colonization
of roots. This finding points to the importance of polar auxin transport inside the
plant when fungal auxin accumulates in plant tissues. This transport and redistribution of auxin is likely to be important in enabling colonization.
Also in cases of LRP emergence in the absence of any fungi, distinct auxin
maxima that establish due to directed auxin transport have been shown to trigger
cell wall remodelling enzymes and to permit cells to separate (Swarup et al. 2008).
A similar mechanism may be involved to permit fungal auxin to activate cell
separation during Hartig net establishment. Due to the lack of identification of
auxin biosynthesis pathways in ECM fungi in contrast to other fungi (Reineke et al.
2008), hitherto no mutants deficient in auxin production are available. Those could
confirm our hypothesis that fungal auxin is crucial for root colonization, but does
not impact LR stimulation.
Interestingly, an antagonist of auxin has also been shown to be involved in ECM
formation. This molecule is the tryptophane-derivative hypaphorine that is produced in large amounts by P. tinctorius upon ECM formation (Beguiristain and
Lapeyrie 1997). It has been proposed that hypaphorine secreted by the fungus
influences root endogenous auxin and thereby regulates symbiosis establishment
(Ditengou et al. 2000; Ditengou and Lapeyrie 2000; Jambois et al. 2005).
Hypaphorine had been shown to interfere with the actin and microtubular cytoskeleton (Ditengou et al. 2003) and with calcium fluxes (Dauphin et al. 2007), at least in
root hairs. However, not all ECM fungi produce hypaphorine. Thus, a general
mechanism based on this compound cannot be proposed. Nonetheless, it cannot
be ruled out as fungi that do not produce hypaphorine may produce other indole
compounds with similar activities.
Signalling in Ectomycorrhizal Symbiosis
135
Fig. 4 Signal exchange during the developing ECM. The hypothetical events in the three phases
of root colonization are related to root developmental changes observed during poplar root
colonization with L. bicolor. During the early phase, root exudates induce hyphae branching and
attract the fungus. The presence of the fungal signal molecules including volatiles stimulates auxin
biosynthesis in plant epidermal cells. This process coincides with LR induction. During the
intermediate phase, when colonization takes place, the fungus releases actively IAA through
PGP-like carriers into the apoplast of root epidermis cell wall. Together with indole compounds
released by the fungus and increased proton effluxes through plasma membrane H+-ATPases. The
induced acidification of the cell wall would facilitate cell spacing and penetration by the fungus.
Cells start to decrease excess IAA through conjugation. In parallel, secreted fungal molecules
including hydrophobines, polysaccharides and small proteins participate to the Hartig net
construction. In the late phase, the fungus still releases IAA into the epidermis cell wall, but
plants import this auxin actively and degrade it to prevent further apoplast acificiation and ongoing
penetration of the fungus. Increased auxin degradation may decrease the total auxin content in the
cells and also influence root growth arrest and root cap decay that were specifically observed in
the late phase. Orange metabolites and proteins are of fungal origin and green ones of plant origin
Taken together, due to its low concentration, fungal auxin is unlikely to be the
trigger of LR stimulation in the early phase of root–ECM fungus interaction.
Sufficient fungal auxin accumulation in plant tissues is probably only possible
136
J. Felten et al.
when the contact of both partners is close enough (mantle/Hartig net). In our model
(Fig. 3), we propose a gradual change of plant and fungal auxin accumulation at the
plant–fungus interface in respective phases of ECM development.
6 Secreted Fungal Molecules as Putative Effectors
Studies developed over the last years have demonstrated that secreted proteins also
act as powerful effectors. The activity of these secreted proteins in the manipulation
of host-cell structure and function has been broadly reported in many
plant–microbial pathosystems (Kamoun 2007). Studies on signalling events in
pathogenicity and symbiosis have highlighted a particular class of effectors
corresponding to secreted small proteins (SSP). Transcript profiling analysis
(Martin et al. 2008) has revealed that these SSPs, so-called Mycorrhiza induced
Small Secreted Proteins (MiSSPs), are specially expressed in the Hartig net infection of the ectomycorhizal symbiont, L. bicolor in poplar.
Identification of the target molecules is crucial for a deeper understanding of the
complex signalling pathways affected in host–symbiont interactions.
7 Evolution of Ectomycorrhizal Signalling
The beginning of ECM fungi evolution is dated to a large time period, probably
about 180–130 mya, and it is possible that they arose at different occasions when
saprotrophic fungi formed symbiotic partnerships (Hibbett et al. 2000; Alexander
2006; Moyersoen 2006). ECM fungi are therefore younger than endomycorrhizal
fungi, which developed 450–350 mya, when plants first settled on land (Simon et al.
1993). The independent evolution of endo- and ectomycorrhizal symbiosis may
account for the difference in their signalling mechanisms.
However, parallels can be drawn between ECM signalling and plant interactions
with nonbeneficial microbes. Different transcriptome studies have revealed that a
defense reaction arises in the plant upon early contact with the fungus, which is
repressed at later stages of colonization (Duplessis et al. 2005; Le Quere et al. 2005;
Sebastiana et al. 2009). Stress/defense responses involve an accumulation of measurable reactive oxygen species (ROS). ROS production during colonization of
roots by ECM fungi is a known phenomenon and is likely to depend on the
compatibility of the interacting partners (Gafur et al. 2004). Commonly during
microbe–plant interactions an innate immunity response triggers the defense reaction in the plant partner. Innate immunity relies on the perception of a microbe/
pathogen associated molecular pattern (MAMP/PAMP) by plant cells via receptors
that activate the defense response (Boller and He 2009). But microbes and
pathogens can release effectors (polysaccharide, proteins, phytotoxins, etc.) that
repress this immune (defense) response. During ECM symbiosis an innate
Signalling in Ectomycorrhizal Symbiosis
137
immunity response has not yet been revealed, but as it is a rather general mechanism, we may suggest that it takes place upon root–ECM fungus interaction. The
fact that during ECM symbiosis establishment the stress response is suppressed
implies that the fungus secretes such an effector.
8 Conclusions
Even though several studies have addressed the specific signals that trigger ECM
symbiosis establishment, the molecules and perception mechanisms involved
remain largely unknown. Unlike bacterial or fungal endosymbiosis, where the
recognition mechanism (ligand/receptor pair) between plant and fungus and parts
of the downstream signalling cascade have been revealed (Stracke et al. 2002;
Gherbi et al. 2008; Olah et al. 2005; D’Haeze and Holsters 2002), the early
signalling in ECM symbiosis is still a black box. Thus, a clear scheme of factors
involved in signalling, recognition, and colonization and their downstream signalling cascade cannot yet be drawn for ECM signalling. However, we propose some
hypothetical events in the context of poplar/Laccaria bicolor interaction (Fig. 4).
We make an attempt to assign them to specific phases in ECM development.
Nonetheless, as data is fragmentary, it is possible that certain molecules are
involved in more phases than the ones we have allocated them to in our models.
References
Aguilar JMM, Shby AM, Richards AJM, Loake GJ, Watson MD, Shaw CH (1988) Chemotaxis of
rhizobium leguminosarum biovar phaseoli towards flavonoid inducers of the symbiotic nodulation genes. J Gen Microbiol 134:2741–2746
Al-Abras K, Bilger I, Martin F, Le Tacon F, Lapeyrie F (1988) Morphological and physiological
changes in ectomycorrhizas of spruce (Picea excelsa (lam.) link) associated with ageing.
New Phytol 110:535–540
Alexander IJ (2006) Ectomycorrhizas – out of Africa? New Phytol 172:589–591
Bari R, Jones JD (2009) Role of plant hormones in plant defence responses. Plant Mol Biol
69:473–488
Bates TR, Lynch JP (2000) Plant growth and phosphorus accumulation of wild type and two root
hair mutants of Arabidopsis thaliana (Brassicaceae). Am J Bot 87:958–963
Beguiristain A, Lapeyrie F (1997) Host plant stimulates hypaphorine accumulation in Pisolithus
tinctorius hyphae during ectomycorrhizal infection while excreted fungal hypaphorine controls
root hair development. New Phytol 136:525–532
Benkova E, Hejatko J (2009) Hormone interactions at the root apical meristem. Plant Mol Biol
69:383–396
Blasius D, Feil W, Kottke I, Oberwinkler F (1986) Hartig net structure and formation in fully
ensheathed ectomycorrhizas. Nord J Bot 6:837–842
Blaudez D, Jacob C, Turnau K, Colpaert JV, Ahonen-Jonnarth U, Finlay R, Botton B, Chalot M
(2000) Differential responses of ectomycorrhizal fungi to heavy metals in vitro. Mycol Res
104:1366–1371
138
J. Felten et al.
Boller T, He SY (2009) Innate immunity in plants: an arms race between pattern recognition
receptors in plants and effectors in microbial pathogens. Science 324:742–744
Buée M, Reich M, Murat C, Morin E, Nilsson RH, Uroz S, Martin F (2009) 454 Pyrosequencing
analyses of forest soils reveal an unexpectedly high fungal diversity. New Phytol 184:449–456
Chalot M, Blaudez D, Brun A (2006) Ammonia: a candidate for nitrogen transfer at the mycorrhizal interface. Trends Plant Sci 11:263–266
Charvet-Candela V, Hitchin S, Ernst D, Sandermann H Jr, Marmeisse R, Gay G (2002) Characterization of an aux/iaa cDNA upregulated in Pinus pinaster roots in response to colonization
by the ectomycorrhizal fungus Hebeloma cylindrosporum. New Phytol 154:769–777
Creelman RA, Mullet JE (1997) Biosynthesis and action of jasmonates in plants. Annu Rev Plant
Physiol Plant Mol Biol 48:355–381
Dauphin A, Gerard J, Lapeyrie F, Legue V (2007) Fungal hypaphorine reduces growth and induces
cytosolic calcium increase in root hairs of eucalyptus globulus. Protoplasma 231:83–88
Dexheimer J, Pargney JC (1991) Comparative anatomy of the host-fungus interface in
mycorrhizas. Experientia 47:312–320
Dexheimer J, Aubert-Dufresne M-P, Gerard J, Le Tacon F, Mousain D (1986) Ultrastructural
localization of acid phosphatase activities in two ectomycorhizas: Pinus nigra nigricans/
Hebeloma crustiliniforme and Pinus pinaster/Pisolithus tinctorius. Lett Botaniq 133:343–352
D’Haeze W, Holsters M (2002) Nod factor structures, responses, and perception during initiation
of nodule development. Glycobiology 12:79R–105R
Ditengou FA, Lapeyrie F (2000) Hypaphorine from the ectomycorrhizal fungus pisolithus
tinctorius counteracts activities of indole-3-acetic acid and ethylene but not synthetic auxins
in eucalypt seedlings. Mol Plant Microbe Interact 13:151–158
Ditengou FA, Beguiristain T, Lapeyrie F (2000) Root hair elongation is inhibited by hypaphorine,
the indole alkaloid from the ectomycorrhizal fungus Pisolithus tinctorius, and restored by
indole-3-acetic acid. Planta 211:722–728
Ditengou FA, Raudaskoski M, Lapeyrie F (2003) Hypaphorine, an indole-3-acetic acid antagonist
delivered by the ectomycorrhizal fungus Pisolithus tinctorius, induces reorganisation of actin
and the microtubule cytoskeleton in Eucalyptus globulus ssp bicostata root hairs. Planta
218:217–225
Duplessis S, Courty PE, Tagu D, Martin F (2005) Transcript patterns associated with
ectomycorrhiza development in eucalyptus globulus and Pisolithus microcarpus. New Phytol
165:599–611
Ek M, Ljungquist PO, Stenstr€
om E (1983) Indole-3-acetic acid production by mycorrhizal fungi
determined by gas chromatography-mass spectrometry. New Phytol 94:401–407
Felten J, Kohler A, Morin E, Bhalerao RP, Palme K, Martin F, Ditengou FA, Legue V (2009) The
ectomycorrhizal fungus Laccaria bicolor stimulates lateral root formation in poplar and
Arabidopsis through auxin transport and signaling. Plant Physiol 151:1991–2005
Felten J, Legue V, Ditengou FA (2010) Lateral root stimulation in the early interaction between
arabidopsis thaliana and the ectomycorrhizal fungus Laccaria bicolor: is fungal auxin the
trigger? Plant Signal Behav 5:864–867
Fries N, Serck-Hanssen K, Dimberg LH, Theander O (1987) Abietic acid, an activator of
Basidiospore germination inectomycorrhizal species of the genus Suillus (Boletaceae). Exp
Mycol 11:360–363
Fukaki H, Tasaka M (2009) Hormone interactions during lateral root formation. Plant Mol Biol
69:437–449
Gafur A, Sch€utzend€
ubel A, Langenfeld-Heyser R, Frizt E, Polle A (2004) Compatible and
incompetent Paxillus involutus isolates for ectomycorrhiza formation in vitro with poplar
(Populus x canescens) differ in H2O2 production. Plant Biol 2004:91–99
Gherbi H, Markmann K, Svistoonoff S, Estevan J, Autran D, Giczey G, Auguy F, Peret B, Laplaze L,
Franche C, Parniske M, Bogusz D (2008) SYMRK defines a common genetic basis for plant root
endosymbioses with arbuscular mycorrhiza fungi, rhizobia, and Frankia bacteria. Proc Natl Acad
Sci USA 105:4928–4932
Signalling in Ectomycorrhizal Symbiosis
139
Gogala N (1991) Regulation of mycorrhizal infection by hormonal factors produced by hosts and
fungi. Experientia 47:331–340
Gomez-Roldan V, Fermas S, Brewer PB, Puech-Pages V, Dun EA, Pillot JP, Letisse F, Matusova R,
Danoun S, Portais JC, Bouwmeester H, Becard G, Beveridge CA, Rameau C, Rochange SF
(2008) Strigolactone inhibition of shoot branching. Nature 455:189–194
Graham JH, Linderman RG (1980) Ethylene production by ectomycorrhizal fungi, Fusarium
oxysporum f. Sp. Pini, and by aseptically synthesized ectomycorrhizae and fusarium-infected
douglas-fir roots. Can J Microbiol 26:1340–1347
Grove M, Spencer G, Rowwedder W, Mandava N, Worley J, Warthen J, Steffens G, FlippenAnderson J, Cook J (1979) Brassinolide, a plant growth-promoting steroid isolated from
Brassica napus pollen. Nature 281:216–217
Gutjahr C, Paszkowski U (2009) Weights in the balance: jasmonic acid and salicylic acid signaling
in root-biotroph interactions. Mol Plant Microbe Interact 22:763–772
Hibbett DS, Gilbert LB, Donoghue MJ (2000) Evolutionary instability of ectomycorrhizal
symbioses in basidiomycetes. Nature 407:506–508
Hilbert JL, Martin F (1988) Regulation of gene expression in ectomycorrhizas. I. Protein changes
and the presence of ectomycorrhiza specific polypeptides in the Pisolithus eucalyptus symbiosis. New Phytol 110:339–346
Hilbert JL, Costa G, Martin F (1991) Ectomycorrhizin synthesis and polypeptide changes during
the early stage of eucalypt mycorrhiza development. Plant Physiol 97:977–984
Ho I (1987a) Comparison of eight Pisolithus tinctorius isolates for growth rate, enzyme activity,
and phytohormone production. Can J For Res 17:31–35
Ho I (1987b) Enzyme activity and phytohormone production of a mycorrhizal fungus, Laccaria
laccata. Can J For Res 17:855–858
Horan DP, Chilvers GA (1990) Chemotropism – the key to ectomycorrhiza formation. New Phytol
116:297–301
Horan DP, Chilvers GA, Lapeyrie F (1988) Time sequence of the infection process in eucalypt
ectomycorrhizas. New Phytol 109:451–458
Ivanchenko MG, Muday GK, Dubrovsky JG (2008) Ethylene-auxin interactions regulate lateral
root initiation and emergence in Arabidopsis thaliana. Plant J 55:335–347
Jacobs PF, Peterson RL, Massicotte G (1989) Altered fungal morphogenesis during early stages of
ectomycorrhiza formation in Eucalyptus pilularis. Scanning Microsc 3:249–255
Jambois A, Dauphin A, Kawano T, Ditengou FA, Bouteau F, Legue V, Lapeyrie F (2005)
Competitive antagonism between IAA and indole alkaloid hypaphorine must contribute to
regulate ontogenesis. Physiol Plant 123:120–129
Jones AR, Kramer EM, Knox K, Swarup R, Bennett MJ, Lazarus CM, Leyser HM, Grierson CS
(2009) Auxin transport through non-hair cells sustains root-hair development. Nat Cell Biol
11:78–84
Kamoun S (2007) (2007) Groovy times: filamentous pathogen effectors revealed. Curr Opin Plant
Biol 187:920–928
Karabaghli-Degron C, Sotta B, Bonnet M, Gay G, Le Tacon F (1998) The auxin transport inhibitor
2,3,5-triiodobenzoic acid (TIBA) inhibits the stimulation of in vitro lateral root formation and
the colonization of the tap-root cortex of norway spruce (Picea abies) seedlings by the
ectomycorrhizal fungus Laccaria bicolor. New Phytol 140:723–733
Kende H, Zeevaart J (1997) The five “classical” plant hormones. Plant Cell 9:1197–1210
Laajanen K, Vuorinen I, Salo V, Juuti J, Raudaskoski M (2007) Cloning of Pinus sylvestris
scarecrow gene and its expression pattern in the pine root system, mycorrhiza and NPA-treated
short roots. New Phytol 175:230–243
Lagrange H, Jay-Allemand C, Lapeyrie F (2001) Rutin, the phenolglycoside from Eucalyptus root
exudates stimulates Pisolithus hyphal growth at picomolor concentrations. New Phytol
150:349–355
Landeweert R, Hoffland E, Finlay RD, Kuyper TW, van Breemen N (2001) Linking plants to
rocks: ectomycorrhizal fungi mobilize nutrients from minerals. Trends Ecol Evol 16:248–254
140
J. Felten et al.
Le Quere A, Wright DP, Soderstrom B, Tunlid A, Johansson T (2005) Global patterns of gene
regulation associated with the development of ectomycorrhiza between birch (Betula pendula
roth.) and Paxillus involutus (batsch) fr. Mol Plant Microbe Interact 18:659–673
Martin F (2007) Fair trade in the underworld: The ectomycorrhizal symbiosis, vol 2, The mycota
biology of the fungal cell. Springer, Heidelberg
Martin F, Nehls U (2009) Harnessing ectomycorrhizal genomics for ecological insights. Curr Opin
Plant Biol 12:508–515
Martin F, Tagu D (1999) Developmental biology of a plant-fungus-symbiosis: the ectomycorrhiza.
In: Varma A, Hock B (eds), Mycorrhiza: structure, function, molecular biology and biotechnology, vol 2. Springer-Verlag berlin, Heidelberg, 51–73
Martin F, Duplessis S, Ditengou F, Lagrange H, Voiblet C, Lapeyrie F (2001) Developmental
cross talking in the ectomycorrhizal symbiosis: signals and communication genes. New Phytol
151:145–154
Martin F, Aerts A, Ahren D, Brun A, Danchin EG, Duchaussoy F, Gibon J, Kohler A, Lindquist E,
Pereda V, Salamov A, Shapiro HJ, Wuyts J, Blaudez D, Buee M, Brokstein P, Canback B,
Cohen D, Courty PE, Coutinho PM, Delaruelle C, Detter JC, Deveau A, DiFazio S, Duplessis S,
Fraissinet-Tachet L, Lucic E, Frey-Klett P, Fourrey C, Feussner I, Gay G, Grimwood J,
Hoegger PJ, Jain P, Kilaru S, Labbe J, Lin YC, Legue V, Le Tacon F, Marmeisse R, Melayah D,
Montanini B, Muratet M, Nehls U, Niculita-Hirzel H, Oudot-Le Secq MP, Peter M,
Quesneville H, Rajashekar B, Reich M, Rouhier N, Schmutz J, Yin T, Chalot M, Henrissat B,
Kues U, Lucas S, Van de Peer Y, Podila GK, Polle A, Pukkila PJ, Richardson PM, Rouze P,
Sanders IR, Stajich JE, Tunlid A, Tuskan G, Grigoriev IV (2008) The genome of laccaria
bicolor provides insights into mycorrhizal symbiosis. Nature 452:88–92
Massicote HB, Perterson RL, Ashford AE (1987) Ontogeny of Eucalyptus piluliris–Pisolithus
tinctorius ectomycorrhizae. I. Light microscopy and scanning electron microscopy. Can J Bot
65:1927–1939
Mensen R, Hager A, Salzer P (1998) Elicitor-induced changes of wall-bound and secreted
peroxidase activities in suspension-cultured spruce (Picea abies) cells are attenuated by auxins.
Physiol Plant 102:539–546
Miersch O, Bohlmann H, Wasternack C (1999) Jasmonates and related compounds from Fusarium
oxysporum. Phytochemistry 50:517–523
Moyersoen B (2006) Pakaraimaea dipterocarpacea is ectomycorrhizal, indicating an ancient
gondwanaland origin for the ectomycorrhizal habit in Dipterocarpaceae. New Phytol
172:753–762
Nylund JE (1980) Symplastic continuity during hartig net formation in Norway spruce
ectomycorrhizas. New Phytol 86:373
Nylund JE, Unestam T (1982) Structure and physiology of ectomycorrhizae I. The process of
mycorrhiza formation in Norway spruce in vitro. New Phytol 91:63–79
Olah B, Briere C, Becard G, Denarie J, Gough C (2005) Nod factors and a diffusible factor from
arbuscular mycorrhizal fungi stimulate lateral root formation in Medicago truncatula via the
DMI1/DMI2 signalling pathway. Plant J 44:195–207
Pitts RJ, Cernac A, Estelle M (1998) Auxin and ethylene promote root hair elongation in
Arabidopsis. Plant J 16:553–560
Plett JM (2010) Ethylene – a key arbitrator to plant–fungal symbiotic interactions? New Phytol
185:868–871
Rahman A, Hosokawa S, Oono Y, Amakawa T, Goto N, Tsurumi S (2002) Auxin and ethylene
response interactions during Arabidopsis root hair development dissected by auxin influx
modulators. Plant Physiol 130:1908–1917
Raudaskoski M, Salo V (2008) Dichotomization of mycorrhizal and NPA-treated short roots in
Pinus sylvestris. Plant Signal Behav 3:113–115
Rayle DL, Cleland RE (1992) The acid growth theory of auxin-induced cell elongation is alive and
well. Plant Physiol 99:1271–1274
Signalling in Ectomycorrhizal Symbiosis
141
Reddy SM, Hitchin S, Melayah D, Pandey AK, Raffier C, Henderson J, Marmeisse R, Gay G
(2006) The auxin-inducible GH3 homologue pp-gh3.16 is downregulated in Pinus pinaster
root systems on ectomycorrhizal symbiosis establishment. New Phytol 170:391–400
Regvar M, Gogala N, Znidarsic N (1997) Jasmonic acid affects mycorrhization of spruce seedlings
with Laccaria laccata. Trees 11:511–514
Reineke G, Heinze B, Schirawski J, Buettner H, Kahmann R, Basse CW (2008) Indole-3-acetic
acid (IAA) biosynthesis in the smut fungus Ustilago maydis and its relevance for increased
IAA levels in infected tissue and host tumour formation. Mol Plant Pathol 9:339–355
Rincon A, Gerard J, Dexheimer J, Le Tacon F (2001) Effect of an auxin transport inhibitor on
aggregation and attachment processes during ectomycorrhiza formation between Laccaria
bicolor S238N and Picea abies. Can J Bot 79:1152–1160
Rincon A, Priha O, Sotta B, Bonnet M, Le Tacon F (2003) Comparative effects of auxin transport
inhibitors on rhizogenesis and mycorrhizal establishment of spruce seedlings inoculated with
Laccaria bicolor. Tree Physiol 23:785–791
Rousseau JVD, Sylvia DM, Fox AJ (1994) Contribution of ectomycorrhiza of the potential
nutrient-absorbing surface of pine. New Phytol 128:639–644
Rudawska ML, Kieliszewska-Rokicka B (1997) Mycorrhizal formation by Paxillus involutus
strains in relation to their IAA-synthesizing activity. New Phytol 137:509–515
Rupp LA, DeVries HE, Mudge KW (1989) Effect of aminocyclopropane carboxylic acid and
aminoethoxyvinylglycine on ethylene production by ectomycorrhizal fungi. Can J Bot
67:483–485
Sebastiana M, Figueiredo A, Acioli B, Sousa L, Pessoa F, Balde A, Pais MS (2009) Identification
of plant genes involved on the initial contact between ectomycorrhizal symbionts (Castanea
sativa – European chestnut and Pisolithus tinctorius). Eur J Soil Biol 45:275–282
Simon L, Bousquet J, Levesque RC, Lalonde M (1993) Origin and diversification of
endomycorrhizal fungi and coincidence with vascular land plants. Natures 363:67–69
Slankis V (1950) Effect of naphthalene acetic acid on dichotomous branching of isolates roots of
Pinus sylvestris. Physiol Plant 3:40–44
Smith SE, Read DJ (2008) Mycorrhizal symbiosis, 3rd edn. Academic, London
Splivallo R, Fischer U, Gobel C, Feussner I, Karlovsky P (2009) Truffles regulate plant root
morphogenesis via the production of auxin and ethylene. Plant Physiol 150:2018–2029
Staswick PE, Serban B, Rowe M, Tiryaki I, Maldonado MT, Maldonado MC, Suza W (2005)
Characterization of an arabidopsis enzyme family that conjugates amino acids to indole-3acetic acid. Plant Cell 17:616–627
Stepanova AN, Yun J, Likhacheva AV, Alonso JM (2007) Multilevel interactions between
ethylene and auxin in Arabidopsis roots. Plant Cell 19:2169–2185
Stracke S, Kistner C, Yoshida S, Mulder L, Sato S, Kaneko T, Tabata S, Sandal N, Stougaard J,
Szczyglowski K, Parniske M (2002) A plant receptor-like kinase required for both bacterial
and fungal symbiosis. Nature 417:959–962
Sun J, Xu Y, Ye S, Jiang H, Chen Q, Liu F, Zhou W, Chen R, Li X, Tietz O, Wu X, Cohen JD,
Palme K, Li C (2009) Arabidopsis asa1 is important for jasmonate-mediated regulation of
auxin biosynthesis and transport during lateral root formation. Plant Cell 21:1495–1511
Swarup K, Benkova E, Swarup R, Casimiro I, Peret B, Yang Y, Parry G, Nielsen E, De Smet I,
Vanneste S, Levesque MP, Carrier D, James N, Calvo V, Ljung K, Kramer E, Roberts R,
Graham N, Marillonnet S, Patel K, Jones JD, Taylor CG, Schachtman DP, May S, Sandberg G,
Benfey P, Friml J, Kerr I, Beeckman T, Laplaze L, Bennett MJ (2008) The auxin influx carrier
LAX3 promotes lateral root emergence. Nat Cell Biol 10:946–954
Tagu D, De Bellis R, Balestrini R, De Vries OMH, Piccoli G, Stocchi V, Bonfante P, Martin F
(2001) Immunolocalization of hydrophobin HYDPt-1 from the ectomycorrhizal basidiomycete
Pisolithus tinctorius during colonization of Eucalyptus globulus roots tinctorius during colonization of Eucalyptus globulus roots. New Phytol 149:127–135
Taylor AFS (2002) Fungal diversity in ectomycorrhizal communities: sample efforts and species
detection. Plant Soil 244:753–760
Terpstra I, Heidstra R (2009) Stem cells: the root of all cells. Semin Cell Dev Biol 20:1089–1096
142
J. Felten et al.
Tranvan H, Habricot Y, Jeannette E, Gay G, Sotta B (2000) Dynamics of symbiotic establishment
between an IAA-overproducing mutant of the ectomycorrhizal fungus Hebeloma
cylindrosporum and Pinus pinaster. Tree Physiol 20:123–129
Umehara M, Hanada A, Yoshida S, Akiyama K, Arite T, Takeda-Kamiya N, Magome H, Kamiya
Y, Shirasu K, Yoneyama K, Kyozuka J, Yamaguchi S (2008) Inhibition of shoot branching by
new terpenoid plant hormones. Nature 455:195–200
Wallender H, Nylund JE, Sundberg B (1992) Ectomycorrhiza and nitrogen effects on root IAA:
result contrary to current theory. Mycorrhiza 1:91–92
Wang JW, Wang LJ, Mao YB, Cai WJ, Xue HW, Chen XY (2005) Control of root cap formation
by microRNA-targeted auxin response factors in Arabidopsis. Plant Cell 17:2204–2216
Wessels JGH (1997) Hydrophobins: proteins that change the nature of the fungal surface. Adv
Microb Physiol 38:1–45
Signalling in the Epichloe¨ festucae: Perennial
Ryegrass Mutualistic Symbiotic Interaction
Carla Eaton, Milena Mitic, and Barry Scott
Abstract Epichloe¨ festucae is a biotrophic fungus that forms symbiotic
associations with temperate grasses of the Festuca and Lolium spp. The association
between E. festucae and L. perenne (perennial ryegrass) is a good experimental
system to study these endophyte–grass associations. Integral to the establishment
and maintenance of these mutualistic associations is mutual communication
between the endophyte and host partner. This communication will likely involve
many well-known signalling pathways. Mitogen-activated protein kinase (MAPK)
cascades and second messenger signalling pathways involving cAMP and calcium
are the main pathways for transduction of signals perceived by cell surface
receptors such as G-protein coupled receptors and histidine kinases. Here we review
what is currently known about signalling mechanisms in the E. festucae–L. perenne
symbiosis including a bioinformatics analysis of the E. festucae genome to identify
which components of these key signalling pathways have been conserved in this
organism. Where pathways are yet to be functionally analysed in E. festucae we
present data from fungal plant pathogens and try to predict possible functions for
these pathways in endophyte associations.
1 Introduction
Epichloe¨ endophytes are a group of sexual and asexual clavicipitaceous fungi
(Clavicipitaceae, Ascomycota) that form symbiotic associations (symbiota) with
temperate grasses of the sub-family Pooideae. There are at least ten different sexual
species (Schardl and Wilkinson 2000) including E. festucae, a natural symbiont of
C. Eaton • M. Mitic • B. Scott (*)
Institute of Molecular BioSciences, Centre for Functional Genomics, Massey University,
Private Bag 11 222, Palmerston North 4442, New Zealand
e-mail: [email protected]; [email protected]; [email protected]
S. Perotto and F. Baluska (eds.), Signaling and Communication in Plant Symbiosis,
Signaling and Communication in Plants 11, DOI 10.1007/978-3-642-20966-6_7,
# Springer-Verlag Berlin Heidelberg 2012
143
144
C. Eaton et al.
Festuca spp. (Leuchtmann et al. 1994). E. festucae also forms a stable mutualistic
association with perennial ryegrass, Lolium perenne, and is proving to be a good
experimental system to study these endophyte–grass associations (Christensen et al.
1997; Scott et al. 2007). The asexual Neotyphodium endophytes are predominantly
interspecific hybrids that form mutualistic associations with their host (Moon et al.
2004).
These biotrophic fungi systemically colonise the intercellular spaces of leaf
primordia, sheaths and blades of vegetative tillers and the inflorescence tissues of
reproductive tillers. During the host reproductive phase, sexual species are capable
of forming external reproductive structures (stromata) that prevent emergence of
the host inflorescence, a disease known as “choke”. Initiation of stroma formation
appears to be triggered by a switch from restricted symbiotic hyphal growth in the
apoplast of the leaf to proliferative pathogenic growth in the outer cell layers and
surface of the leaf.
In these mutualistic associations, the major benefits to the fungal symbiont are
access to nutrients from the host apoplast and a means of dissemination through the
seed. Benefits to the host include increased tolerance to both biotic (e.g. insect and
mammalian herbivory) and abiotic stresses (e.g. drought).
2 Biotrophic Lifestyle
Both N. lolii and E. festucae form stable, asymptomatic mutualistic associations
with L. perenne and are vertically transmitted through the seed following
colonisation of the ovule (Philipson and Christey 1986). After fertilisation, the
endophyte colonises the embryo sac and as the seed matures hyphae spreads
throughout the embryo (Philipson and Christey 1986; May et al. 2008). During
seed germination, hyphae colonise the shoot apical meristem (SAM) from where
they enter leaf primordia and axillary buds, the meristematic zones from which new
shoots develop. Hyphae appear to grow within the intercellular spaces of the shoot
apex by polarised tip growth, and are frequently branched to form a reticulated
mycelial network throughout the shoot apex. After colonisation and attachment to
the cells within the leaf expansion zone hyphae further elongate by intercalary
division and extension (Christensen et al. 2008). This novel pattern of growth
allows the fungal endophyte to synchronise its growth with that of the grass host
and avoid mechanical shear as the leaf cells are displaced away from the leaf
expansion zone. Hyphae grow parallel to the leaf axis but occasionally branch
and fuse with adjacent hyphae to establish an extensive intercellular network within
the leaf (Fig. 1). At all stages of leaf growth the hyphae are firmly attached to the
mesophyll cell walls, providing an ideal connection for nutrient transport and
intercellular signalling between host and symbiont (Fig. 2).
Signalling in the Epichloe¨ festucae: Perennial Ryegrass Mutualistic Symbiotic Interaction
145
Fig. 1 Symbiotic phenotype of E. festucae noxR mutant. Confocal depth series images (1 mm) of
longitudinal sections through perennial ryegrass (Lolium perenne) leaves infected with wild type
(WT) and noxR mutant stained with Alexafluor (WGA-AF488) and aniline blue. The image shows
hyphae (fluorescent green) of Epichloe¨ festucae growing in close association with plant cells.
Strongly illuminated points indicate hyphal septa
Fig. 2 Transmission electron micrograph showing an E. festucae hypha growing in the intercellular space between two perennial ryegrass (L. perenne) cells. Bar ¼ 1 mm
146
C. Eaton et al.
3 Regulation of Hyphal Growth in Culture and In Planta
The highly restricted pattern of epichloë endophyte growth in planta implies there
must be signalling between the host and the fungus in order to maintain this tightly
regulated mutualistic association. Fungal genes involved in host signalling have
been identified using a forward genetic screen in which plasmid insertional mutagenesis was used to generate mutants which were screened for any disruption in
their association with perennial ryegrass (Tanaka et al. 2006). This strategy
identified a mutant that induced severe stunting of leaf growth and resulted in
premature senescence of the host. Mutant hyphae were hyperbranched in the leaves
and were no longer aligned parallel to the leaf axis. In addition, fungal biomass in
planta was significantly greater than in wild-type associations. This dramatic plant
interaction phenotype was caused by disruption of noxA, a gene that encodes the
catalytic subunit of the NADPH oxidase (Nox) complex.
3.1
Regulation of Fungal Growth In Planta
by the Nox Complex
The multi-subunit Nox complex is most well characterised in mammalian systems
where its role in the respiratory (oxidative) burst response of phagocytes to invading pathogens is well established (Lambeth 2004). The enzymatic function of this
complex is to catalyse the conversion of molecular oxygen to the reactive oxygen
species (ROS) superoxide, via an NADPH-dependent reaction in which electrons
are transferred from intracellular NADPH via FAD and two heme cofactors to
molecular oxygen, generating superoxide on the extracellular face of the plasma
membrane (Sumimoto et al. 2005).
E. festucae NoxA is most similar to the mammalian Nox2 isoform, known as
gp91phox. In mammalian phagocytes, gp91phox is embedded in the plasma membrane where it associates with p22phox to form the flavocytochrome b558 complex,
the catalytic core of the oxidase (Groemping and Rittinger 2005). In response to an
activating signal, the small GTPase Rac1, and a heterotrimer consisting of p67phox
(the Nox activator), p47phox (the Nox organiser) and p40phox translocate to the
plasma membrane and associate with gp91phox, leading to the production of
superoxide (Groemping and Rittinger 2005). To date only homologues of gp91phox
(NoxA and NoxB), p67phox (NoxR) and Rac1 (RacA) have been identified in fungi
(Takemoto et al. 2007) Homologues of p40phox and p47phox appear to be absent in
fungal genomes, suggesting that fungal specific proteins are required for the
recruitment of NoxR and RacA to the plasma membrane (Takemoto et al. 2006).
Recent work showed that the yeast polarity proteins Bem1 and Cdc24 interact with
NoxR and preferentially localise to actively growing hyphal tips and septa,
suggesting they may play an analogous role in fungi to p40phox and p47phox in
mammals (Takemoto et al. 2011). Importantly, plants infected with E. festucae DnoxR
Signalling in the Epichloe¨ festucae: Perennial Ryegrass Mutualistic Symbiotic Interaction
147
(Takemoto et al. 2006), DracA (Tanaka et al. 2008) or DbemA (Takemoto et al. 2011)
mutants also exhibit a defective symbiotic interaction, demonstrating that these
additional components of the Nox complex are also essential for symbiotic maintenance. However, it is unclear whether ROS produced by the fungal Nox complex
signals to the fungus itself, to the host plant or to both partners. If the fungal Nox
complex is localised to the plasma membrane and has the same membrane topology
as the phagocytic Nox complex (Sumimoto et al. 2005), ROS would be released
into the intercellular space (apoplast) of the host or into the media for cultures
grown ex planta. Formation of electron dense cerium perhydroxide deposits on the
plasma membrane and cell wall of hyphae in planta provides support for the
hypothesis that ROS is released into the apoplast (Fig. 3) (Tanaka et al. 2008).
This ROS would have the potential to either signal to the plant and/or activate
endophyte membrane channels to transduce a signal to the fungal cytoplasm.
However, examination of fungal ROS production in culture using nitroblue tetrazolium (NBT), which reacts with superoxide anions to form a dark blue formazan
precipitate, revealed the presence of a basipetal ROS gradient within hyphal tips
(Tanaka et al. 2006; Semighini and Harris 2008). This suggests that ROS is being
generated internally either from Nox located on internal membranes or from
alternative enzyme sources (Tanaka et al. 2006; Semighini and Harris 2008; Grissa
et al. 2010). Further research is required on the cellular location and topology of
fungal ROS producing enzymes to resolve this issue.
3.2
ROS as Signalling Molecules
Given the importance of the Nox complex for symbiotic maintenance, the question
arises: how do ROS produced by this complex regulate fungal growth in planta and
maintain these associations? ROS are well suited to a role as signalling molecules
as they are small and highly diffusible, can be rapidly produced by an NADPH
oxidase in response to an activating stimulus and rapidly broken down by antioxidant enzymes (Hancock et al. 2001; Lalucque and Silar 2003). In addition, as Nox
Fig. 3 ROS production by
E. festucae constitutively
active RacA strain in
perennial ryegrass.
Transmission electron
micrograph of H2O2
localisation in meristematic
tissue. Cerium chloride
deposits are indicated by
arrowheads. fc fungal cell,
pc plant cell
148
C. Eaton et al.
enzymes are NADPH dependent, their activity is directly linked to cellular metabolism (Lalucque and Silar 2003). However, the main problem with ROS as signalling molecules is their lack of target specificity. As their name suggests, ROS are
highly reactive. Hydroxyl radicals, for example, will react with any nearby organic
molecule, making them unlikely signalling molecules (Baier et al. 2005). Hydrogen
peroxide and superoxide are less reactive so could function as signalling molecules
(Baier et al. 2005), although they are unlikely to have specific receptors due to their
simple chemical structure (Neill et al. 2002). Instead, it is likely that ROS signal
indirectly by oxidation of key signalling proteins, in a reversible process known as
redox regulation. Changes in the oxidation state of proteins often lead to changes in
their activity (Buchanan and Balmer 2005). The basic mechanism of oxidation of
proteins by hydrogen peroxide is well characterised. The hydrogen peroxide molecule reacts with thiolate cysteine residues (RS), converting them into sulfenate
ions (RSO), which rapidly react with nearby thiol cysteine residues (RSH) to form
a disulfide bond (Forman et al. 2004). This may lead to conformational changes in
the protein that can either reduce or enhance activity, affect cofactor binding or
affect binding to other proteins (Allen and Tresini 2000). This oxidation is reversible via antioxidants such as glutathione or thioredoxin, which reduce the oxidised
cysteine residue (Lambeth 2004). Two of the best-characterised targets of redox
regulation are the transcription factors OxyR (from E. coli) and Yap1 (from
Saccharomyces cerevisiae) (Delaunay et al. 2002; Kim et al. 2002). Reduced
OxyR is able to bind DNA, but cannot activate transcription. Oxidation by hydrogen peroxide induces formation of a disulfide bond, which enables OxyR to activate
transcription (Kim et al. 2002). In contrast, Yap1 is not directly oxidised by
hydrogen peroxide. Instead, a glutathione peroxidase, Gpx3, is oxidised by hydrogen peroxide and forms an intermolecular disulfide bond to Yap1. This bond is then
converted to an intramolecular disulfide bond within Yap1, which activates Yap1
and prevents its export from the nucleus (Delaunay et al. 2002). However, it is
important to note that the ROS involved in oxidation of these proteins is not
necessarily produced by a Nox complex, as this family of enzymes is not found
in unicellular organisms such as yeast (Lalucque and Silar 2003). Indeed, most ROS
are produced as by-products of aerobic respiration. However, ROS can also be
generated enzymatically by some peroxidases (Chen and Schopfer 1999) and
potentially by ferric reductases (Takeda et al. 2007; Semighini and Harris 2008).
A range of other enzymatic and non-enzymatic proteins are also subject to redox
regulation. These include the protein tyrosine phosphatases, which in contrast to
OxyR and Yap1, are strongly inhibited by oxidation (Leslie et al. 2004). The
activity of ion channels is also regulated by ROS signalling (Baier et al. 2005).
For example, ROS increases outflow of potassium ions through a potassium channel
known as human ether-a-gogo (HERG) (Taglialatela et al. 1997), and hydrogen
peroxide activates calcium channels in the plasma membrane of Arabidopsis
thaliana guard cells (Pei et al. 2000). A number of translation initiation and
elongation factors, and ribosomal and RNA binding proteins have also been
found to be targets of ROS oxidation, providing a link between ROS and translation
(Buchanan and Balmer 2005).
Signalling in the Epichloe¨ festucae: Perennial Ryegrass Mutualistic Symbiotic Interaction
149
ROS molecules are also known to regulate important cellular signalling pathways,
and in particular there is a strong link between ROS and calcium (Ca2+) signalling in a
variety of organisms. Interactions between Ca2+ and ROS signalling systems are very
complex and may be both stimulatory and inhibitory, depending on the type of target
proteins, the ROS species, dose, duration of exposure and the cell context (Mori and
Schroeder 2004; Yan et al. 2006). Many ROS generation processes are calcium
dependent. Some NOX enzymes have Ca2+ binding regulatory domains and are
directly regulated by calcium, as is the case for mammalian NOX5 (Jagnandan et al.
2007), DUOX1 and DUOX2 isoforms (Lambeth 2004). All NOX enzymes in plants
are directly regulated by calcium (Torres et al. 1998). In the social amoeba
Dictyostelium discoideum, signalling of NOXA and NOXB, homologues of
gp91phox, is regulated by calcium through Alg-2B, a calcium binding protein, while
NOXC has its own calcium binding domains (Lardy et al. 2005). However, deletion of
pef-1, the N. crassa homologue of alg-2b, was unable to suppress either the nox-1 or
nox-2 developmental phenotypes (Cano-Domı́nguez et al. 2008). NOXC isoforms
present in some fungi also contain calcium-binding domains (Takemoto et al. 2007).
Calcium signals themselves may be regulated by the redox state of the cell, as is the
case for the aforementioned ROS-activated calcium channels of the A. thaliana guard
cells. ROS also regulates calcium uptake in A. thaliana growing root cells. A. thaliana
rhd2 mutants are defective in calcium uptake and their root development is severely
impaired, resulting in stunted roots and short root hairs. The rhd2 gene encodes an
NADPH oxidase (Nox) and ROS produced by this protein accumulates in wild-type
growing root hairs but are absent from the rhd2 mutant cells. A direct link between
ROS and calcium signalling was established when it was shown that addition of ROS
to rhd2 mutant cells partially suppress the rhd2 phenotype and stimulates the activity
of plasma membrane hyperpolarisation-activated Ca2+ channels (Foreman et al. 2003;
Takeda et al. 2008). The connection between ROS and Ca2+ signalling in E. festucae
and its importance for the regulation of symbiosis remain to be established.
3.3
Additional Regulators of Fungal Growth In Planta
In addition to components of the fungal Nox complex, two other E. festucae genes
have also been shown to be essential for symbiotic maintenance. The first of these is
the stress-activated MAP kinase, sakA, homologous to mammalian p38MAPK or
S. cerevisiae Hog1. Deletion of E. festucae sakA had a dramatic effect on the
fungal–host interaction phenotype (Eaton et al. 2010). Infected plants were stunted
and senesced within 2–3 months after planting. There was also a dramatic change in
plant development, with formation of bulb-like structures at the base of the tillers
and almost complete loss of the photoprotective anthocyanin pigments. Fungal
growth in planta was deregulated, with hyphal hyperbranching, increased fungal
biomass and colonisation of host vascular bundles. Using high-throughput mRNA
sequencing to compare fungal and plant gene expression between a wild-type
association and the DsakA mutant association revealed a number of changes
150
C. Eaton et al.
consistent with a transition from restricted hyphal growth in the wild-type mutualistic association to proliferative growth in the mutant association.
Deletion of E. festucae sidN, encoding an iron siderophore, also leads to
disruption of the E. festucae–perennial ryegrass symbiotic interaction (Johnson
et al. 2007). The product of SidN is proposed to be a novel ferrichrome-like
extracellular siderophore (Johnson 2008). The breakdown of the symbiosis in plants
infected with this mutant highlights the importance of iron homeostasis, particularly the ability of the endophyte to access iron from the apoplast, to maintain a
stable symbiotic interaction.
3.4
Regulation of Fungal Growth by Calcium Signalling
A central dogma of fungal biology is that hyphae grow by tip growth, a situation
analogous to growth of plant pollen tubes and root hairs. Calcium is widely
recognised as a crucial molecule in signalling pathways regulating polarised
growth. In plants, calcium ions play a crucial role in root hair growth and elongation
of pollen tubes. The tip of the growing pollen tube has a cytosolic Ca2+ gradient,
which is essential for tube growth and is formed by an influx of extracellular Ca2+
through stretch-activated ion channels. In the pollen protoplast, these channels are
found at the site of tube germination and only after incubation in germination
medium (Feijo et al. 2001; Dutta and Robinson 2004).
While tip-localised Ca2+ gradients have been observed in polarised fungal
hyphal growth there is some debate about the in vivo significance of these gradients
because of the lack of reliable techniques for the measurement of cytosolic Ca2+
concentrations. Some studies suggest that a Ca2+ gradient at the hyphal tip, formed
by Ca2+ release from the endoplasmic reticulum (ER) in response to activation of
stretch-activated phospholipase C and generation of IP3, is essential for hyphal
growth (Silverman-Gavrila and Lew 2002, 2003). However, IP3-regulated Ca2+
channels are yet to be identified in fungi. Additional studies are needed in order to
fully understand the role of Ca2+ gradients at the hyphal tip in regulating growth.
However, the role of calcium signalling pathways in regulating hyphal growth and
branching is well documented (Jackson and Heath 1993; Torralba and Heath 2001).
In N. crassa, disturbance of the intracellular calcium concentration causes
hyperbranching of the growing tip (Schmid and Harold 1988) and application of
high concentrations of exogenous calcium restores a wild-type growth pattern to
hyperbranched mutants (Kawano and Said 2005). Furthermore, disruption of
calcineurin, a Ca2+-regulated phosphatase, induced hyperbranching in N. crassa,
dissipation of the tip-high Ca2+ gradient and eventually growth arrest, suggesting
calcineurin is essential for normal tip growth (Prokisch et al. 1997). While nothing
is known so far about the importance of Ca2+ gradients in the regulation of hyphal
growth of E. festucae, genes encoding homologues of calcium signalling genes
identified in other fungi are found in the E. festucae genome (Nguyen et al. 2008)
(Fig. 4).
Signalling in the Epichloe¨ festucae: Perennial Ryegrass Mutualistic Symbiotic Interaction
151
Fig. 4 Proposed components of Ca2+ signalling pathways in E. festucae. Calcium ions enter the
cytoplasm from the extracellular spaces and intracellular depots through a system of Ca2+ channels
and are actively transported out of the cytoplasm through various Ca2+ pumps and exchangers.
In the cytoplasm, Ca2+ regulates activity of various enzymes like phospholipase C and
phosphatases. E. festucae contains three Ca2+ channels, various Ca2+ pumps and transporters,
four PLCs, three Ca2+/calmodulin regulated kinases, one or two copies of calcineurin A and
various other calcium regulated proteins
4 Signalling Pathways
Undoubtedly, some of the most important signal transduction pathways in filamentous fungi are MAP kinase, heterotrimeric G protein and two-component regulatory
pathways. However, with the exception of the stress-activated MAP kinase, these
pathways are yet to be studied in epichloë endophytes or any other mutualistic plant
symbiotic fungi. This section, therefore, provides an overview of the roles these
pathways play in pathogenesis by phytopathogenic fungi. Results of a bioinformatic
152
C. Eaton et al.
survey of the E. festucae Fl1 (E894) and E2368 genomes to identify which
components of these pathways are encoded in these genomes will also be presented,
allowing for predictions as to what roles these pathways may play in
endophyte–grass associations.
4.1
MAP Kinase Pathways
Mitogen-activated protein (MAP) kinase pathways are perhaps some of the most
conserved and crucial signalling pathways found in eukaryotes. These pathways
transduce diverse physical or chemical signals detected at the cell surface into
changes in gene expression via a phosphorylation cascade consisting of three highly
conserved protein kinases – the MAP kinase kinase kinase (MAPKKK), MAP
kinase kinase (MAPKK) and MAP kinase (MAPK). Filamentous fungi generally
possess three MAP kinase pathways, which respond to different stimuli – the stressactivated MAP kinase pathway, pheromone response/filamentation pathway and
cell wall integrity pathway. The mechanism by which these pathways signal has
been extensively reviewed by others (Banuett 1998; Gustin et al. 1998), so will not
be covered here.
4.1.1
The Stress-Activated MAP Kinase Pathway
As the name suggests, the stress-activated MAP kinase pathway is activated in
response to perception of an environmental stress. The best characterised of these
pathways is the Hog1 (high osmolarity glycerol) pathway of S. cerevisiae. In
S. cerevisiae, this pathway is primarily involved in the response to osmotic stress,
with mutants in this pathway unable to grow under conditions of high osmolarity
(Brewster et al. 1993). In contrast, the homologous Sty1/Spc1 (suppressor of
tyrosine phosphatase/suppressor of phosphatase 2C) pathway of the model fission
yeast Schizosaccharomyces pombe responds to a variety of different stresses
including osmotic, oxidative, temperature and UV stresses (Millar et al. 1995;
Shiozaki and Russell 1995). In this regard, the stress-activated MAP kinase
pathways of filamentous fungi are more similar to that of S. pombe, being able to
respond to multiple environmental stresses (reviewed in Rispail et al. 2009).
Interestingly, these pathways in filamentous fungi have been adapted to not only
transmit stress signals, but to enhance the virulence of pathogenic fungi. For
example, in the dimorphic fungus Mycosphaerella graminicola, the causal agent
of Septoria tritici leaf blotch on wheat, MgHog1 mutants are non-pathogenic due to
an inability to switch to filamentous growth required for infection (Mehrabi et al.
2006). Similarly, Cochliobolus heterostrophus hog1 mutants have reduced disease
symptoms on maize (Igbaria et al. 2008), and Botrytis cinerea Dbcsak1 MAPK
mutants display reduced virulence on bean plants (Segm€uller et al. 2007). However,
Signalling in the Epichloe¨ festucae: Perennial Ryegrass Mutualistic Symbiotic Interaction
153
not all filamentous fungal stress-activated MAP kinases are involved in virulence,
as Magnaporthe grisea Dosm1 mutants (Dixon et al. 1999), Bipolaris oryzae Dsrm1
mutants (Moriwaki et al. 2006) and Colletotrichum lagenarium osc1 mutants
(Kojima et al. 2004) are still fully pathogenic. As discussed in the previous section,
signalling through the stress-activated MAP kinase, SakA, is required for maintenance of a mutualistic interaction between E. festucae and perennial ryegrass
(Eaton et al. 2010). This raises an interesting conundrum – why are stress-activated
MAP kinase mutants of some phytopathogenic fungi less virulent, whereas the
E. festucae DsakA mutant has increased virulence? It would be interesting to see
whether other components of the E. festucae stress-activated MAP kinase pathway
are also necessary for symbiotic maintenance, or whether regulation of the symbiosis by SakA occurs independently of the MAP kinase pathway. In phytopathogenic
fungi, other components of the stress-activated MAP kinase pathway are often also
required for virulence, including homologues of the membrane sensors Sho1 and
Msb2 (Lanver et al. 2010), the small GTPase Cdc42 (Zheng et al. 2009), p21activated kinases Cla4 and Ste20 (Smith et al. 2004; Rolke and Tudzynski 2008)
and the MAPKKK Ste11 (Zhao et al. 2005). All additional components of the
E. festucae stress-activated MAP kinase pathway found in the genomes of Fl1 and
E2368 are presented in Fig. 5 and Table 1.
4.1.2
The Pheromone Response/Filamentation Pathway
In yeast, the pheromone response/filamentation pathway is primarily involved in
perception of a pheromone, which binds to a receptor on the cell surface, activating
a heterotrimeric G protein signalling pathway (discussed in more detail later),
ultimately leading to activation of the MAP kinase pathway (reviewed in Rispail
et al. 2009). The activated Fus3 MAP kinase then targets effectors that induce cell
cycle arrest, polarised growth and formation of shmoo cells required for mating
(Rispail et al. 2009). Similar to the stress-activated MAP kinase pathway, the
pheromone response/filamentation pathway has also been adapted to a role in
virulence of phytopathogenic fungi. Indeed, this pathway has been shown to be
required for virulence in nearly all phytopathogenic fungi examined, including
M. grisea (Xu and Hamer 1996), Fusarium oxysporum (Di Pietro et al. 2001,
2003), C. heterostrophus (Lev et al. 1999), Alternaria alternata (Lin et al. 2010),
Botrytis cinerea (Zheng et al. 2000), C. lagenarium (Takano et al. 2000) and
Claviceps purpurea (Mey et al. 2002). C. purpurea, a pathogen of rye, is very
closely related to the epichloë endophytes therefore we predict that the pheromone
response/filamentation pathway will also play a role in symbiosis between epichloë
endophytes and their host plants. Homologues of all major components of the
pheromone response/filamentation MAP kinase pathway have been identified in
the E. festucae Fl1 and E2368 genomes, and are presented in Fig. 6 and Table 1.
Interestingly, no obvious homologue of the yeast cyclin-dependent kinase inhibitor
Far1 could be identified in the E. festucae genomes. Far1 homologues have been
identified in other filamentous fungi, including F. graminearum, M. grisea and
154
C. Eaton et al.
Fig. 5 Proposed components of the stress-activated MAP kinase pathway in E. festucae. Schematic showing components of the stress-activated MAP kinase pathway encoded in the genome of
E. festucae Fl1 and E2368. Proteins are coloured as follows: orange, membrane sensor; red,
phosphatase; blue, kinase of the MAPK cascade; pink, transcription factor; light green, guanine
nucleotide exchange factor (GEF); cyan, unspecified kinase; purple, p21-activated kinase; dark
green, other function. Dashed grey boxes indicate proteins not found encoded in the E. festucae
genomes. Where proteins have been characterised in other filamentous fungi those gene names are
used, otherwise naming is based on S. cerevisiae homologues. Adapted from Rispail et al. (2009)
N. crassa (Rispail et al. 2009). The apparent absence of a Far1 homologue suggests
that it was either lost in E. festucae or it has diverged to such an extent that it is no
longer identifiable as a homologue.
Signalling in the Epichloe¨ festucae: Perennial Ryegrass Mutualistic Symbiotic Interaction
155
Table 1 E. festucae genes were identified by tBlastn analysis of the E. festucae Fl1 genome
(http://lims.ca.uky.edu/CLS894Blast/blast/blast.html) with M. grisea protein sequences
M. grisea
E value
%
Comment
E. festucae
S. cerevisiae
locus
identity
gene name
gene namea
mck1
Bck1
MGG_00883 0
45
MAP kinase kinase kinase
bemA
Bem1
MGG_01702 4e-84
58
SH3-domain protein
mkk1
MGG_06482 0
64
MAP kinase kinase
pkc1
MGG_08689 0
70
Protein kinase C
mps1
Mpk1
MGG_04943 e-177
80
MAP kinase
msg5
MGG_15140 0
52
MAPK phosphatase
pakB
Ste20
MGG_12821 0
59
p21-activated kinase
sakA
Hog1
MGG_01822 e-154
94
MAP kinase
pbs2
MGG_10268 e-166
53
MAP kinase kinase
pmk1
Fus3
MGG_09565 e-179
97
MAP kinase
ptp2
MGG_00912 3e-38
28
Tyrosine protein
phosphatase
ptp3
MGG_01376 0
39
Tyrosine protein
phosphatase
ssk2/22
MGG_00183 0
60
MAP kinase kinase kinase
ste11
MGG_12855 0
56
MAP kinase kinase kinase
ste50
MGG_05199 e-127
72
Protein kinase regulator
ste7
MGG_00800 e-106
67
MAP kinase kinase
pre1
Ste2
MGG_06452 8e-55
30
Pheromone receptor
pre2
Ste3
MGG_04711 3e-52
34
Pheromone receptor
rgsA
Sst2
MGG_14517 e-140
52
Regulator of G protein
signalling
sepA
Bni1
MGG_04061 0
65
Formin
ste12
MGG_12958 0
75
Transcription factor
cdc42
MGG_00466 1e-91
91
Rho GTPase
cdc24
MGG_09697 0
50
Guanine nucleotide
exchange factor
gnb1
Ste4
MGG_05201 0
89
Guanine nucleotide-binding
protein b subunit
gng1
Ste18
MGG_10193 8e-39
87
Guanine nucleotide-binding
protein g subunit
gna1
Gpa1
MGG_00365 e-160
98
Guanine nucleotide-binding
protein a subunit
pakA
Cla4
MGG_06320 0
73
p21-activated kinase
ssk1
MGG_02897 2e-78
40
Cytoplasmic response
regulator
hpt1
Ypd1
MGG_07173 9e-28
32
Histidine phosphotransfer
protein
sln1
MGG_07312 0
26
Osmosensor histidine kinase
sho1
MGG_09125 e-103
65
Transmembrane
osmosensor
msb2
MGG_06033 6e-50
31
Mucin family member
ptc1
MGG_05207 e-157
44
Protein phosphatase 2C
homolog
ptc2
MGG_01351 4e-45
61
(continued)
156
C. Eaton et al.
Table 1 (continued)
E. festucae
S. cerevisiae
gene name
gene namea
M. grisea
locus
E value
%
Comment
identity
Protein phosphatase 2C
homolog
atfA
Sko1
MGG_08212 e-111
58
bZIP transcription factor
rck1/2
MGG_08547 e-160
68
Serine-threonine protein
kinase
msn2/4
MGG_00501 1e-66
41
Zinc finger transcription
factor
rlmA
Rlm1
MGG_01204 7e-47
28
MADS-box transcription
factor
mcm1
MGG_02773 2e-28
70
MADS-box transcription
factor
wsc1
MGG_04325 4e-37
40
Plasma membrane sensor
wsc2/3
MGG_03761 0
42
Plasma membrane sensor
rom1/2
MGG_03064 0
67
Guanine nucleotide
exchange factor
tus1
MGG_12644 0
47
Guanine nucleotide
exchange factor
sac7
MGG_06390 e-149
41
GTPase activating protein
rho1
MGG_07176 2e-53
90
GTPase activating protein
sit4
MGG_03911 e-107
81
Serine-threonine
phosphatase
mbp1/swi4
MGG_08463 e-169
57
DNA-binding component of
the SBF complex
swi6
MGG_09869 0
51
Transcription cofactor
ppz1/2
MGG_00149 0
72
Protein phosphatase Z
E values for this analysis and the amino acid sequence identity between the M. grisea protein and
the predicted polypeptide sequence for the E. festucae homologue are given. Proteins chosen for
this analysis were based on the MAP kinase pathway schemes presented in Dean et al. (2005) and
Rispail et al. (2009). E. festucae gene models are available on requestaWhere the E. festucae genes
are named differently to their S. cerevisiae homologues, the S. cerevisiae names are also given
4.1.3
The Cell Wall Integrity Pathway
In fungi, the cell wall integrity pathway acts to mediate cell wall remodelling and
biogenesis in response to cell wall stress, and during certain stages of the cell cycle
(Levin 2005). It does this through a variety of means, including regulation of the
glucan synthase and reorganisation of the actin cytoskeleton (Levin 2005). In
phytopathogenic fungi, this pathway has also been adapted for a role in virulence.
For example, the C. heterostrophus mps1 MAPK mutant displays reduced virulence
(Igbaria et al. 2008), the C. lagenarium maf1 MAPK mutant is defective in
appressorium development and so displays reduced pathogenicity (Kojima et al.
2002), the M. grisea mps1 mutant is non-pathogenic (Xu et al. 1998), and
F. graminearum MGV1 MAPK is required for full virulence (Hou et al. 2002).
The C. purpurea MAPK, CPMK2, is also essential for virulence (Mey et al. 2002),
Signalling in the Epichloe¨ festucae: Perennial Ryegrass Mutualistic Symbiotic Interaction
157
Fig. 6 Proposed components of the pheromone response/filamentation MAP kinase pathway in
E. festucae. Schematic showing components of the pheromone response/filamentation MAP
kinase pathway encoded in the genome of E. festucae Fl1 and E2368. Proteins are coloured as
follows: orange, G-protein coupled receptor; red, phosphatase; blue, kinase of the MAPK cascade;
pink, transcription factor; light green, guanine nucleotide exchange factor (GEF); purple, p21activated kinase; yellow, GTPase; dark green, other function. Dashed grey boxes indicate proteins
not found encoded in the E. festucae genomes. Where proteins have been characterised in other
filamentous fungi those gene names are used, otherwise naming is based on S. cerevisiae
homologues. Adapted from Rispail et al. (2009)
and given its close phylogenetic relationship to E. festucae we would predict that
the cell integrity MAP kinase of E. festucae will be required for symbiotic maintenance. It is likely that other components of this pathway will also be required for
158
C. Eaton et al.
symbiosis signalling, given their importance for virulence in phytopathogenic fungi
(Jeon et al. 2008). All components of the E. festucae Fl1 and E2368 cell integrity
MAP kinase pathway have been identified and are presented in Fig. 7 and Table 1.
Fig. 7 Proposed components of the cell wall integrity MAP kinase pathway in E. festucae.
Schematic showing components of the cell wall integrity MAP kinase pathway encoded in the
genome of E. festucae Fl1 and E2368. Proteins are coloured as follows: orange, membrane sensor;
red, phosphatase; blue, kinase of the MAPK cascade; pink, transcription factor; light green,
guanine nucleotide exchange factor (GEF) or GTPase activating protein (GAP); yellow, GTPase;
cyan, unspecified kinase. Dashed grey boxes indicate proteins not found encoded in the E. festucae
genomes. Where proteins have been characterised in other filamentous fungi those gene names are
used, otherwise naming is based on S. cerevisiae homologues. Adapted from Rispail et al. (2009)
Signalling in the Epichloe¨ festucae: Perennial Ryegrass Mutualistic Symbiotic Interaction
4.2
159
Heterotrimeric G-Protein Signalling
Filamentous fungal heterotrimeric G proteins were first identified in N. crassa
(Turner and Borkovich 1993), and much of what we know about G-protein signalling has been determined in this organism (reviewed in Li et al. 2007). In
unstimulated conditions, a G-protein coupled receptor (GPCR) on the cell surface
is bound to a heterotrimer of Ga (GDP bound), Gb and Gg proteins. Binding of an
extracellular ligand to the GPCR triggers the GDP on the Ga subunit to be
exchanged for GTP. This induces release of the heterotrimer, which then
dissociates into GTP-bound Ga and a Gb/Gg dimer. These components are then
able to interact with downstream effectors. Hydrolysis of the Ga-bound GTP back
to GDP induces the Ga to reassociate with the Gb/Gg dimer and the GPCR at the
membrane (reviewed in Li et al. 2007). The involvement of the heterotrimeric G
proteins in virulence of phytopathogenic fungi has been extensively studied.
Components of the heterotrimeric G protein signalling pathway encoded in the
E. festucae Fl1 and E2368 genomes are presented in Table 2.
4.2.1
Ga Subunits
Most filamentous fungi contain three Ga subunits, which tend to play different roles
in the cell (Li et al. 2007). In M. grisea, only one of these three Ga genes, magB, is
required for virulence (Liu and Dean 1997). In C. heterostrophus only CGA1 is
Table 2 E. festucae genes were identified by tBlastn analysis of the E. festucae Fl1 genome
(http://lims.ca.uky.edu/CLS894Blast/blast/blast.html) with N. crassa protein sequences
E. festucae N. crassa locus E value % identity Comment
gene name
gna1
NCU06493
e-157
97
Guanine nucleotide-binding protein a subunit
gna2
NCU06729
1e-98
77
Guanine nucleotide-binding protein a subunit
gna3
NCU05206
1e-75
76
Guanine nucleotide-binding protein a subunit
gnb1
NCU00440
e-165
85
Guanine nucleotide-binding protein b subunit
gng1
NCU00041
2e-24
90
Guanine nucleotide-binding protein g subunit
orp1
NCU01735
2e-38
32
G-protein coupled receptor
pre1
NCU00138
1e-50
20
G-protein coupled receptor
pre2
NCU05758
8e-41
25
G-protein coupled receptor
gpr1a
NCU00786
6e-53
22
G-protein coupled receptor
gpr1b
NCU00786
3e-9
27
G-protein coupled receptor
gpr2
NCU04626
5e-93
40
G-protein coupled receptor
gpr4
NCU06312
6e-38
24
G-protein coupled receptor
gpr5
NCU00300
1e-41
34
G-protein coupled receptor
gpr6
NCU09195
4e-49
51
G-protein coupled receptor
E values for this analysis and the amino acid sequence identity between the N. crassa protein and
the predicted polypeptide sequence for the E. festucae homologue are given. Proteins chosen for
this analysis were based on those present in the N. crassa genome, as presented in Li et al. (2007).
As this analysis focused on identification of homologues of the N. crassa G protein signalling
proteins, there may be additional G-protein coupled receptors encoded in the E. festucae genome
that were not identified by this analysis. E. festucae gene models are available on request
160
C. Eaton et al.
required for normal appressorium formation (Horwitz et al. 1999), and in
C. parasitica only cpg-1 is required for virulence (Gao and Nuss 1996). In contrast,
both Gpa1 and Gpa3 of Mycosphaerella graminicola are required for pathogenicity, while Gpa2 is dispensable (Mehrabi et al. 2009). Similarly, both bcg1 and bcg2
of B. cinerea are required for normal virulence (Gronover et al. 2001). Both
Fusarium oxysporum fga1 (Jain et al. 2002) and Stagonospora nodorum gna1
(Solomon et al. 2004) are required for pathogenicity, but it is still unclear whether
the other Ga genes from these fungi are also involved in pathogenicity. Interestingly, Ustilago maydis has four Ga genes, but only one of these, gpa3, is required
for pathogenesis (Regenfelder et al. 1997). Similar to most other filamentous fungi,
the E. festucae genomes encode three Ga proteins (Table 2). Given the key role
these gene products play in phytopathogenesis, it is very likely that at least one of
these proteins will be required for symbiotic maintenance.
4.2.2
Gb and Gg Subunits
In contrast to the multiple Ga genes possessed by most filamentous fungi, generally
only one gene for Gb and one for Gg are present in fungal genomes (Li et al. 2007).
The Gb gene product, in particular, has a key role in phytopathogenicity. For
example, C. heterostrophus CGB1 is required for normal virulence on maize
(Ganem et al. 2004), and F. oxysporum fgb1 is required for pathogenicity on
cucumber (Jain et al. 2003). In addition, M. grisea mgb1 mutants are non-pathogenic (Nishimura et al. 2003), and M. graminicola (Mehrabi et al. 2009) and
C. parasitica (Kasahara and Nuss 1997) Gb mutants have reduced virulence. It is
unclear yet what role the Gg subunit may play in virulence of phytopathogenic
fungi. The E. festucae Fl1 and E2368 genomes encode products for one Gb and one
Gg, which are highly conserved with homologues from other fungi (Table 2). We
predict that the E. festucae Gb subunit will be crucial for symbiotic maintenance.
4.2.3
G-Protein Coupled Receptors (GPCRs)
GPCRs are the membrane bound component of the G-protein signalling pathway
that respond to changes in the environment or ligand binding and transmit the
response to the heterotrimeric G proteins. In N. crassa, these proteins have diverse
functions including pheromone response, cAMP response and carbon sensing
(reviewed in Li et al. 2007). At present there are no reports that we are aware of
on the role of GPCRs in virulence of phytopathogenic fungi. We predict that the
products of at least some of these genes will be crucial for transmitting metabolic
signals from the host to the symbiont. The E. festucae genomes appear to contain
genes for at least nine GPCRs, as identified by their homology to the ten GPCRs
originally identified in N. crassa (which is now known to encode considerably more
GPCRs than this) (Table 2). Interestingly, the E. festucae genomes do not appear to
encode a homologue of the N. crassa opsin NOP-1. In addition, no homologue of
Signalling in the Epichloe¨ festucae: Perennial Ryegrass Mutualistic Symbiotic Interaction
161
N. crassa GPR-3 could be identified, but two genes showing high homology to
GPR-1 were identified.
4.3
Two-Component Regulatory Signalling
Two-component regulatory system (TCS) pathways comprise a membrane-bound
histidine kinase and a cytosolic response regulator which, in response to signal
perception by the histidine kinase is autophosphorylated, resulting in activation of
downstream effectors. The importance of these pathways in environmental sensing
by prokaryotes is well established (reviewed in Chang and Stewart 1998). A fungal
TCS pathway was first identified in yeast, and its role upstream of the HOG1 stressactivated MAP kinase pathway is now well established (reviewed in Saito and
Tatebayashi 2004). In filamentous fungi, a TCS was first characterised in N. crassa,
following the identification of the two-component histidine kinase, NIK-1 (OS-1)
(Alex et al. 1996). With the sequencing of the N. crassa genome, ten additional HKs
were identified (Galagan et al. 2003). Interestingly, N. crassa contains only two
response regulators (RRs), RRG-1 (Jones et al. 2007) and RRG-2 (Banno et al.
2007), suggesting that signals from all 11 HKs must be channelled through the two
RRs. By comparison to most prokaryotic TCSs, fungal TCSs are generally more
complex with the HK being comprised of both a histidine kinase domain and a
response regulator receiver domain. Hence, they are referred to as hybrid histidine
kinases (HHKs). The histidine kinase domain is autophosphorylated in response to
an activating signal and the phosphate transferred to the RR receiver domain within
the HHK. In another difference to most prokaryotic systems, the phosphate on the
HHK is first transferred to a histidine phosphotransfer protein (HPT) protein, which
in turn transfers the phosphate to the RR, resulting in activation of downstream
effectors (Stock et al. 2000; Banno et al. 2007).
4.3.1
Hybrid Histidine Kinase
In phytopathogenic fungi, TCS pathways have been adapted for a role in virulence.
The HHKs BOS1 from B. cinerea (Viaud et al. 2006), Fhk1 from F. oxysporum
(Rispail and Di Pietro 2010) and CpHK2 from C. purpurea (Nathues et al. 2007)
have all been shown to be necessary for normal virulence. In contrast, M. grisea
Hik1 is not required for pathogenicity (Motoyama et al. 2005). The genomes of
E. festucae Fl1 and E2368 contain at least nine HHKs, based on a comparison of the
11 genes in N. crassa (Fig. 8 and Table 3). Interestingly, only one phytochrome is
encoded by the E. festucae genome compared to the two, PHY-1 and PHY-2,
encoded by the N. crassa genome.
162
C. Eaton et al.
Fig. 8 Proposed components of two component signalling pathways in E. festucae. Schematic
showing components of two component signalling pathways encoded in the genome of E. festucae
Fl1 and E2368. Hybrid histidine kinases (HHKs) are represented in orange. HHKs were identified
based on homology to the 11 HHKs encoded in the N. crassa genome. Additional novel HHKs
may also be encoded in the E. festucae genomes. HPt histidine phosphotransferase, RR response
regulator
Table 3 E. festucae genes were identified by tBlastn analysis of the E. festucae Fl1 genome
(http://lims.ca.uky.edu/CLS894Blast/blast/blast.html) with N. crassa protein sequences
E. festucae gene name N. crassa locus E value % identity Comment
rrgA
NCU01895
e-142
33
Response regulator
rrgB
NCU02413
1e-75
40
Response regulator
hpt1
NCU01489
3e-24
39
Histidine phosphotransfer protein
os1
NCU02815
0
79
Histidine kinase
phy1
NCU04834
0
41
Histidine kinase
tcsA
NCU00939
0
45
Histidine kinase
fphA
NCU01823
0
56
Histidine kinase
chk1
NCU01833
0
53
Histidine kinase
luxQ
NCU02057
e-168
49
Histidine kinase
sln1
NCU04615
0
27
Histidine kinase
hhk8
NCU00939
2e-99
39
Histidine kinase
hhk9
NCU09520
e-100
22
Histidine kinase
E values for this analysis and the amino acid sequence identity between the N. crassa protein and
the predicted polypeptide sequence for the E. festucae homologue are given. Proteins chosen for
this analysis were based on those present in the N. crassa genome (Galagan et al. 2003). As this
analysis focused on identification of homologues of the N. crassa two-component signalling
proteins, there may be additional histidine kinases encoded in the E. festucae genome that were
not identified by this analysis. E. festucae gene models are available on request
Signalling in the Epichloe¨ festucae: Perennial Ryegrass Mutualistic Symbiotic Interaction
4.3.2
163
Response Regulator and Histidine Phosphotransfer Protein
Very little information is available on the role of fungal response regulators in
phytopathogenesis. However, in Magnaporthe oryzae one RR (SSK1) and one RRlike gene (RIM15) were shown to be required for virulence (Motoyama et al. 2008).
In addition, in C. heterostrophus SSK1 but not SKN7 is required for virulence, and in
Gibberella zeae, ssk1 mutants show reduced pathogenicity (Oide et al. 2010). The
genomes of E. festucae Fl1 and E2368 encode two response regulators, homologous
to N. crassa genes for RRG-1 and RRG-2 (Fig. 8 and Table 3). Given the precedence
for a role of RRs in virulence of the phytopathogenic fungi it is likely that at least one
of the two E. festucae RRs will be required for symbiotic maintenance. To date there
are no reports of a role for the HPt in phytopathogenesis, possibly because mutants of
this gene are lethal as found for A. nidulans (Vargas-Perez et al. 2007). E. festucae
contains one HPt gene, and like in A. nidulans it is predicted to be essential.
4.4
Second Messenger Signalling Pathways
In order to sense and respond to internal and external environmental signals, cells
have developed signalling systems that transmit signals from cell surface receptors
to their intracellular targets using second messengers. Second messengers are small
molecules whose cytoplasmic concentration is controlled in response to extracellular/intracellular signals. This section reviews two of the most important fungal
second messengers: cyclic adenosine monophosphate (cAMP) and calcium.
4.4.1
cAMP Signalling
cAMP regulates a wide range of developmental, metabolic and pathogenic processes in fungi. However, despite regulating a diverse range of processes,
components of cAMP signalling are highly conserved (D’Souza and Heitman
2001). Synthesis of cAMP is catalysed by adenylate cyclase and its degradation
is facilitated by phosphodiesterases. The activity of these enzymes is regulated by
various extracellular signals. Adenylate cyclase is directly regulated by
heterotrimeric G-protein signalling through protein kinase A (PKA), a key enzyme
involved in most cAMP regulated processes. In its inactive state, PKA is a tetramer
composed of two regulatory and two catalytic subunits. When the cytoplasmic
concentration of cAMP increases, cAMP binds to the regulatory subunits causing
disassociation of the catalytic subunits. The two catalytic subunits act downstream
on various transcription factors and metabolic enzymes.
In filamentous fungi, cAMP has a key role in the regulation of growth and
developmental processes. Inactivation of the PKA regulatory subunit disrupts
polarised growth in N. crassa (Bruno et al. 1996) and Aspergillus niger (Saudohar
et al. 2002). cAMP is also important for regulating plant infection by fungal
164
C. Eaton et al.
pathogens, likely due to a role in regulating morphological switching which is often
required for pathogenesis. For example, in U. maydis cAMP is required for formation of the dikaryon and the subsequent morphological switch to filamentous
growth required for infection (reviewed in Kahmann et al. 1999). Mutations in
adenylate cyclase, uac1, or the catalytic subunit of PKA, adr1, induce filamentous
growth of haploid cells while mutation in the PKA regulatory subunit, ubc1, as
expected, has the opposite effect and results in a multiple budding phenotype (Gold
et al. 1994; Durrenberger et al. 1998). Both adr1 and ubc1 are also involved in the
second stage of the infection process, with adr1 mutants being non-pathogenic and
ubc1 mutants able to penetrate the host but unable to induce tumours. These results
suggest cAMP is required for the penetration process but inhibits later steps in
infection, such as tumour development. In the maize pathogen Fusarium
verticillioides, disruption of the PKA catalytic subunit CPK1 or adenylate cyclase
FAC1 leads to defects in radial growth, but only fac1 mutants show significantly
reduced virulence, suggesting there may be another copy of the catalytic subunit
gene which regulates the infection process (Choi and Xu 2010). In M. graminicola,
disruption of either the regulatory or catalytic PKA subunit significantly reduces
virulence (Mehrabi and Kema 2006). Disruption of adenylate cyclase in B. cinerea
disrupts sporulation and reduces the rate of lesion development during the infection
process (Klimpel et al. 2002). cAMP signalling has diverse roles in M. grisea.
Deletion of the PKA catalytic subunit, CPKA, dramatically reduces appressorium
formation (Mitchell and Dean 1995). Deletion of the adenylate cyclase gene,
MAC1, also reduces appressorium formation but it also affects vegetative growth,
conidiation and conidial germination (Choi and Dean 1997). Mutations in the PKA
regulatory subunit (SUM1) suppressed the growth and development phenotypes of
mac1, including appressorium formation (Adachi and Hamer 1998). Under noninducing conditions SUM1 mutants undergo precocious development of
appressoria, suggesting CPKA is not the only catalytic subunit present in M. grisea.
Homologues of adenylate cyclase and the PKA catalytic and regulatory subunits are
found in the genomes of E. festucae Fl1 and E2368. Disruption of acyA (adenylate
cyclase) in E. festucae reduced radial growth and increased conidiation (Voisey
et al. 2007). The most pronounced effects in planta were a reduction in infection
rate and an increase in hyphal branching, although host plants did not become
stunted as was observed for noxA and racA mutants.
4.4.2
Ca2+ as a Second Messenger
Calcium is a universal secondary messenger, involved in a broad range of processes
from gene expression to apoptosis (Bootman et al. 2001), and is proposed to be the
most versatile biological messenger known (Cheng and Lederer 2008). Cytoplasmic calcium concentration is maintained at a very low level (100–500 nM) compared with that in the extracellular spaces and internal stores (1–5 mM) (Halachmi
and Eilam 1989; Permyakov and Kretsinger 2009). A calcium signal is generated when
the cytoplasmic Ca2+ concentration increases, which in turn activates Ca2+-regulated
Signalling in the Epichloe¨ festucae: Perennial Ryegrass Mutualistic Symbiotic Interaction
165
effectors. Calcium ions are transported from the cytoplasm by calcium pumps and
ion exchangers.
Calcium Signalling in Fungi
Fungal calcium signalling has been best characterised in the budding yeast
S. cerevisiae. While relatively little is known about calcium signalling in filamentous fungi, it appears to be more complex than in S. cerevisiae (Zelter et al. 2004).
This section will provide an overview of the main components of fungal Ca2+
signalling pathways, including a bioinformatic survey of calcium signalling
components present in the E. festucae genome (Table 4 and Fig. 4). Calcium
signalling proteins are highly conserved across the fungal kingdom, but with
some variation in copy number. While few of the identified calcium signalling
genes have been functionally characterised in E. festucae, systematic functional
analysis of Ca2+ genes in the phytopathogenic fungus M. oryzae using RNA
silencing provides an insight into the importance of these pathways for plant–fungal
interactions (Nguyen et al. 2008).
Calcium Channels and Transporters
Three Ca2+ permeable channels have been characterised in S. cerevisiae: the
voltage-gated channel Cch1p, stretch-activated Mid1p channel and the transient
receptor potential-like Ca2+ channel Yvc1p. Disruption of mid1 induces loss of cell
viability after pheromone-induced cell differentiation, with the mutant displaying
low Ca2+ uptake. This loss of viability can be restored by incubation in a high
extracellular Ca2+ concentration medium (Iida et al. 1994). Dcch1 mutants appear
identical to Dmid1 mutants, suggesting Cch1p and Mid1p are components of the
same Ca2+ permeable channel (Paidhungat and Garrett 1997). The Yvc1p channel is
located in the vacuolar membrane (Palmer et al. 2001), and is involved in the
calcium response triggered by hyperosmotic shock (Denis and Cyert 2002; Zhou
et al. 2003). Knock-down of M. oryzae Cch1p leads to reduced sporulation and
appressorium formation but the strains remain pathogenic (Nguyen et al. 2008).
Knock-down of Mid1p resulted in a similar phenotype, whereas knock-down of
Yvc1p strongly affected conidia and appressoria formation (Nguyen et al. 2008).
Interestingly, deletion of C. purpurea mid1 resulted in a complete loss of pathogenicity and production of appressoria-like structures not normally seen in
C. purpurea (Bormann and Tudzynski 2009). Like other filamentous fungi,
E. festucae contains homologues of the Ca2+ channel proteins Cch1, Mid1 and
Yvc1 (Table 4). Given the close relationship between E. festucae and C. purpurea it
is likely that E. festucae mid1 may be involved in regulating symbiosis.
The role of Ca2+ ATPases (Ca2+ pumps) and exchangers is to maintain low
cytoplasmic Ca2+ concentrations by actively removing Ca2+ from the cytoplasm. In
fungi, these proteins work together with the phosphatase calcineurin in regulating
166
C. Eaton et al.
Table 4 E. festucae genes were identified by tBlastn analysis of the E. festucae Fl1 genome
(http://lims.ca.uky.edu/CLS894Blast/blast/blast.html) with M. oryzae protein sequences
Gene
M. oryzae
E
%
Comment
name
locus
value Identity
cch1
MGG_5643 0
52
Ca2+ channel
mid1
MGG_12128 0
29
Ca2+ channel
yvc1
MGG_9228 0
59
Ca2+ channel
cnaA1 MGG_07456 0
85
Calcineurin catalytic subunit; present in Fl1 and E2368
cnaA2 MGG_07456 0
62
Calcineurin catalytic subunit; present in E2368, absent
from Fl1
cnaB
MGG_06933 0
64
Calcineurin regulatory subunit
Cam
MGG_06884 0
100
Calmodulin
ncs1
MGG_01550 0
83
Frequenin
alg-2
MGG_04818 0
78
Penta-EF-hand Ca2+ binding protein
cmkA
MGG_09912 0
65
Ca2+/Calmodulin-dependent protein kinase
cmkB
MGG_00925 0
73
Ca2+/Calmodulin-dependent protein kinase
cmkC
MGG_06421 0
43
Ca2+/Calmodulin-dependent protein kinase
plc1
MGG_02444 0
44
Phospholipase C
plc2
MGG_00211 0
54
Phospholipase C
plc3
MGG_15018 0
34
Phospholipase C
plc4
MGG_08315 0
42
Phospholipase C
MGG_05332 0
46
pmcA1 MGG_02487 0
59
Calcium-transporting ATPase
pmcA2 MGG_07971 0
49
Calcium-transporting ATPase
pmcA3 MGG_04890 0
49
Calcium-transporting ATPase
pmr1
MGG_11727 0
70
Secretory pathway Ca2+-ATPase
pmr2
MGG_04550 0
81
Secretory pathway Ca2+-ATPase
cat1
MGG_13279 0
52
K/Na-ATPase
cat2
MGG_05078 0
69
Probably K/Na-ATPase
cat3
MGG_12005 0
75
Unknown pump specificity
cat4
MGG_06295 0
58
Unknown pump specificity
exc1
MGG_01638 0
41
Na+/Ca2+-exchanger
exc2
MGG_11454 0
52
Vacuolar Ca+ transporter
exc3
MGG_08710 0
27
Vacuolar Ca+ transporter
exc4
MGG_04159 0
54
Vacuolar Ca+ transporter
exc5
MGG_01381 0
53
Calcium permease
hyp1
MGG_06475 0
64
Hypothetical protein; Could be related to Ca2+ signalling
anxA7 MGG_06847 0
63
Annexin A7
hyp2
MGG_00654 0
48
Hypothetical protein; EF hand domain-containing
protein
ndeh1 MGG_04140 0
66
NADH dehydrogenase; Could be related to Ca2+
signalling
sal1
MGG_01072 0
53
Ca2+-binding mitochondrial carrier
hyp3
MGG_02342 0
49
Hypothetical protein; Could be related to Ca2+ signalling
ara1
MGG_07066 0
71
Ca2+-binding mitochondrial carrier protein
sagA
MGG_06180 0
76
Contains EF hands
cdc4
MGG_09470 0
58
Myosin regulatory light chain cdc4
ndeh2 MGG_06276 0
65
NADH dehydrogenase; Could be related to Ca2+
signalling
(continued)
Signalling in the Epichloe¨ festucae: Perennial Ryegrass Mutualistic Symbiotic Interaction
167
Table 4 (continued)
Gene
M. oryzae
E
%
Comment
name
locus
value Identity
dun1
MGG_01596 0
58
DNA damage-response kinase
rck2
MGG_08547 0
69
Ca2+/Calmodulin-dependent protein kinase-like protein
hyp4
MGG_03548 0
30
Putative tropomiosin-1 alpha chain
cnx
MGG_01607 0
72
Calnexin
hyp5
MGG_08154 0
46
Calmodulin like protein
kin4
MGG_01196 0
47
Could be related to calcium signalling
E values for the BLASTP analysis and the amino acid sequence identity between the M. oryzae
protein and the predicted polypeptide sequence for the E. festucae homologue are given. Proteins
chosen for this analysis were based on those present in the M. oryzae genome (Nguyen et al. 2008).
Other calcium signalling genes may be present in the E. festucae. E. festucae gene models are
available on request
the cellular response to stress induced by high salt concentration (Cunningham and
Fink 1996; Park et al. 2001). PMR1 homologues in A. niger and Aspergillus
fumigatus are required for maintaining Ca2+ homeostasis in these fungi (Yang
et al. 2001; Pinchai et al. 2010). However, A. fumigatus pmrA, encoding a Ca2+
ATPase, is not required for pathogenicity (Pinchai et al. 2010). E. festucae contains
three PMC1 (vacuolar Ca2+ ATPases) and two PMR1 (Golgi Ca2+ ATPases)
homologues, and five Ca2+ exchangers (Table 4).
Phospholipase C
In S. cerevisiae, deletion of PLC1 was shown to have pleiotropic effects, including
variable rates of survival depending on the genetic background (Yoko-o et al.
1993). Mutants display sensitivity to osmotic stress, temperature stress and show
defects in the utilisation of carbon sources other than glucose (Flick and Thorner
1993). PLC1 works together with the Gpr1p/Gpa2p GPCR to transduce glucoseinduced calcium signals (Flick and Thorner 1993; Tisi et al. 2002). In M. oryzae,
disruption of plc1 resulted in reduced pathogenicity, presumably due to disturbance
of calcium fluxes essential for the infection process (Rho et al. 2009). E. festucae
contains four PLC homologues (Table 4). Given the pathogenicity defects of PLC1
mutants of M. oryzae the homologue from E. festucae may be important for
signalling and maintenance of the symbiosis.
Calmodulin
The calcium sensor calmodulin is involved in many cell processes through the
regulation of targets such as the Ca2+/calmodulin-dependent protein phosphatase,
calcineurin or Ca2+/calmodulin dependent protein kinases. The S. cerevisiae calmodulin gene, CMD1, is essential for viability (Davis et al. 1986). While little is
168
C. Eaton et al.
known about the role calmodulin plays in phytopathogenicity, expression of
M. oryzae calmodulin is upregulated during appressorium formation (Liu and
Kolattukudy 1999). Additionally, calmodulin preferentially localises to germ
tubes and appressoria, suggestive of a role in the infection process (Ma et al.
2009). In support of this, silencing of calmodulin was found to dramatically reduce
pathogenicity of M. oryzae. E. festucae contains one, highly conserved CAM
homologue (Table 4).
Ca2+/Calmodulin-Dependent Protein Kinases
An important group of calcium-regulated enzymes is the Ca2+/calmodulin-dependent protein kinases (CaMKs). CaMKs are serine/threonine protein kinases with an
amino terminal catalytic domain and a carboxy-terminal regulatory domain. The
regulatory domain consists of an autoinhibitory region and the Ca2+/calmodulin
(Ca2+/CaM) binding region. The autoinhibitory domain regulates the activity of the
enzyme by interacting with the catalytic domain to prevent substrate binding or
distort the catalytic site. Binding of Ca2+ activated calmodulin with the calmodulinbinding domain results in a conformational change that releases the catalytic
domain to overcome this autoinhibition. After the initial activation, the enzyme
may remain active independently of the presence of Ca2+/CaM. Furthermore, the
enzyme may require additional modifications, such as phosphorylation, for full
activation. Fungal CaMKs play important roles in fungal development as well as
interactions with other organisms. S. cerevisiae contains two CaM kinases, CMK1
and CMK2 (Ohya et al. 1991). These are involved in thermotolerance (Iida et al.
1995) and cell survival following pheromone-induced cell cycle arrest (Moser et al.
1996). In the filamentous fungus A. nidulans, three Ca2+/CaM-dependent protein
kinases, CmkA, CmkB and CmkC, have been reported. CmkA and CmkB are bona
fide CaMKs, whereas CmkC is a CaMKK, which phosphorylates CmkB in vitro.
Disruption of cmkA or cmkB is lethal, while expression of a constitutively active
form of CmkA prevents spores from entering the first nuclear division cycle
(Dayton and Means 1996; Dayton et al. 1997; Joseph and Means 2000). In
N. crassa, deletion of CAMK-1 induces a transient slow growth phenotype (Yang
et al. 2001). CaM kinases also appear to play a role in phytopathogenicity. M.
oryzae cmk1 mutants have delayed spore germination and appressorium formation,
leading to a reduced ability to infect host plants (Liu et al. 2009). Additionally,
inhibitors of CaMK block the Colletotrichum gloeosporioides infection process
(Kim et al. 1998). However, not all CaM kinases are essential for phytopathogenicity, as loss of any of the three CaMKs of Stagnospora nodorum does not
prevent pathogenicity (Solomon et al. 2006). E. festucae contains homologues of A.
nidulans cmkA, cmkB and cmkC (Table 4). However, none of these genes appear to
be required for establishment or maintenance of the symbiotic interaction between
E. festucae and perennial ryegrass (Mitic 2011).
Signalling in the Epichloe¨ festucae: Perennial Ryegrass Mutualistic Symbiotic Interaction
169
Calcineurin
Calcineurin is a Ca2+/calmodulin-dependent protein phosphatase (PP2B) comprised
of a catalytic subunit (CnA) and a regulatory subunit (CnB). The catalytic subunit is
a polypeptide with four different domains: catalytic domain, CnB binding domain,
Ca2+/CaM binding domain and an autoinhibitory domain. As for CaMKs, the
autoinhibitory domain inhibits enzyme activity and this inhibition is relieved by
Ca2+/CaM binding (Rusnak and Mertz 2000). Calcineurin signalling, which occurs
via the calcineurin-responsive transcription factor CRZ1, is important for regulation
of ion stress, cell wall integrity, hyphal growth and a number of other developmental
processes (Stie and Fox 2008). Calcineurin signalling is also important for many
fungal–plant associations. In U. maydis, calcineurin is essential for virulence, with
mutants unable to form tumours (Egan et al. 2009). In M. oryzae, treatment with the
calcineurin inhibitor cyclosporin A blocks appressorium formation, and RNAi
silencing of the calcineurin A gene, MCNA, significantly reduces appressorium
formation (Choi et al. 2009b). Disruption of M. oryzae CRZ1 confirmed that
calcineurin signalling is required for the infection process, as mutants seldom
developed infectious hyphae (Choi et al. 2009a), possibly due to the reduced turgor
pressure of the mutant compared to wild type (Zhang et al. 2009). B. cinerea crz1
mutants are defective in penetration, but this can be rescued by addition of Mg2+
(Schumacher et al. 2008). An analysis of the genome sequences of E. festucae
strains E2368 and E984 (Fl1) identified two copies of cnaA in the former and just
one in the latter (Table 4). A cnaA deletion mutant of Fl1 induced a severe HR-like
response when seedlings of perennial ryegrass were infected with this mutant (Mitic
2011), suggesting calcineurin signalling does play a role in symbiotic maintenance.
5 Host Defence Response
Plants recognise potentially pathogenic microorganisms, such as bacteria and fungi,
and react to their presence by activation of a multi-layered host defence mechanism.
Activation of the first layer of the plant immune response depends on recognition of
pathogen-associated molecular patterns (PAMPs). Perception of PAMPs by the
plant activates PAMP-triggered immunity (PTI), which, among other responses,
induces rapid ion flux across the membrane, ROS production, changes in gene
expression and cell wall reinforcement (Zipfel 2008). Many pathogens have
evolved various effector proteins that can interfere with PTI, thereby allowing the
infection process to continue. NBS-LRR proteins are key for plant recognition of
microbial effector proteins. This recognition is species specific and results in
effector-triggered immunity (ETI). ETI usually involves host cell death that is
triggered by a hypersensitive response (Jones and Dangl 2006). Chitin is the best
studied fungal PAMP. Suppression of a gene encoding a chitin receptor in rice,
CEBiP, leads to a reduction in the oxidative burst response and expression of genes
that normally define a plant chitin-induced response (Kaku et al. 2006).
170
C. Eaton et al.
Symbiotic fungi, such as endophytes and mycorrhizal fungi need to avoid host
defences in order to establish a functional symbiosis. However, it remains unclear
how this avoidance is achieved in different fungi. Based on a PTI/ETI host response
model, avoidance of host defences could be achieved by masking of fungal PAMPs
or through suppression of host immune responses triggered by fungal effectors. It is
also possible that the plant itself prevents a defence response upon specific recognition of the symbiotic partner. Indeed, transcriptional analysis of the association
between Pinus sylvestris and the ectomycorrhizal fungus Laccaria bicolor revealed
an initial upregulation of some defence and cell wall biosynthesis related genes,
followed by a downregulation of these genes in the later stages of mycorrhizal
establishment (Heller et al. 2008).
In epichloë endophyte associations it remains unclear how the fungus evades any
host defence response – whether the fungus is able to somehow “hide” from the host,
or whether the host recognises the fungus but does not mount a response. While
wild-type E. festucae does not induce a host defence response some E. festucae
symbiotic mutants that form pathogenic associations with perennial ryegrass do
activate host defences. High throughput mRNA sequencing of perennial ryegrass
associations with E. festucae-wild-type and a MAP kinase mutant strain (DsakA),
revealed a dramatic up-regulation of host defence-related genes including pathogenicity response (PR) genes and NBS-LRR genes (Eaton et al. 2010). It is unclear
what changes to the DsakA mutant induced these host defence processes. A possible
explanation for the host defence response seen in plants infected with the DsakA
mutant is a change in the fungal cell wall composition or structure that allows
detection by the host. Although the composition of the cell walls of wild-type
E. festucae and the DsakA mutant have yet to be investigated, transmission electron
micrographs show that the cell walls of the DsakA mutant hyphae in the host
pseudostem often appear much thinner, and more diffuse than those of wild type,
possibly indicating a change in cell wall composition (Eaton et al. 2010).
6 Signalling Associated with Synthesis
of Bioprotective Metabolites
Four main classes of biologically active metabolites have been identified in grass
hosts infected with E. festucae: peramine, indole diterpenes (principally lolitrem B),
ergot alkaloids (principally ergovaline) and lolines. Peramine is a potent feeding
deterrent of adult Argentine stem weevil (ASW; Listronotus bonariensis), an economically important pest of L. perenne in New Zealand agricultural ecosystems.
Lolitrem B also has biological activity against some insect larvae but is better known
as the causative agent of “ryegrass staggers”, a neuromuscular disorder that affects
animals grazing on pastures with high toxin levels. Although the fitness benefits of
ergot alkaloids are still to be defined, they are implicated in herbivory protection
against some insects and nematodes. However, ergot alkaloids are best known for
Signalling in the Epichloe¨ festucae: Perennial Ryegrass Mutualistic Symbiotic Interaction
171
the livestock grazing disorder, “fescue toxicosis”. Loline alkaloids have potent antiinsecticidal properties and can accumulate to very high levels in grass tissues.
Genes for the synthesis of peramine (Tanaka et al. 2005), indole diterpenes
(Young et al. 2005, 2006), ergot alkaloids (Panaccione et al. 2001; Wang et al.
2004; Fleetwood et al. 2007) and lolines (Spiering et al. 2005) have been cloned and
the biochemical function of the protein products inferred from a combination of
chemical and genetic analysis and by a comparison with the known functions of
homologues in other filamentous fungi. With the exception of peramine, which
appears to require just a single gene product, PerA, for its synthesis, the other
classes of metabolites require multiple gene products, encoded by sets of genes that
are organised in distinct clusters within the E. festucae genome. At least ten genes
are required for lolitrem (ltm) and related indole diterpene biosynthesis (Young
et al. 2006), ten genes are required for ergot alkaloid biosynthesis (eas) (Schardl
et al. 2006; Fleetwood et al. 2007) and nine genes are required for loline (lol)
(Spiering et al. 2005) biosynthesis. Using the E. festucae–perennial ryegrass symbiotic association as a model experimental system we have shown that the per, eas
and ltm genes are all preferentially and highly expressed in planta, but not expressed
in culture, even in the presence of crude plant extracts (Tanaka et al. 2005; Young
et al. 2005, 2006; Fleetwood et al. 2007; May et al. 2008). When a promoter of the
ltmM (PltmM) gene was fused to the gusA reporter, GusA activity was detected in
most vegetative and reproductive tissues, including the developing seed, but not in
axenic E. festucae cultures (May et al. 2008). Taken together these results provide
strong evidence that plant-specific signalling is required for expression of all three
biosynthetic pathways. With the exception of N. uncinatum, lolines have not been
detected in axenic cultures of epichloë endophytes (Blankenship et al. 2001).
However, even in N. uncinatum-containing symbiota the lol genes are more highly
expressed in planta compared to culture, confirming that these genes are also
symbiotically regulated.
A distinctive structural feature of these E. festucae secondary metabolite clusters
is their association with a complex array of Type I (RNA based) and Type II (DNA
based) transposon relics, sequences known to promote a chromatin state that is
transcriptionally silent (Martienssen and Colot 2001). Recent work has shown that
chromatin remodelling is required for activation of secondary metabolite biosynthesis in Aspergillus spp. (Shwab et al. 2007; Bok et al. 2009; Reyes-Dominguez et al.
2010). Therefore, chromatin remodelling is predicted to be an important component
of E. festucae secondary metabolite gene activation in planta. How this occurs in
response to plant host signals is a key question to understanding the symbiosis.
7 Future Challenges
A key challenge in the future will be identifying plant metabolites and their
associated receptors which sense and transduce these signals to activate (1) ROS
expression and (2) secondary metabolite biosynthesis, in order to control hyphal
172
C. Eaton et al.
growth and regulate synthesis of bioprotective molecules to maintain a mutually
beneficial symbiotic interaction. The crucial cellular zone for interaction between
symbiont and host is the apoplast. It is therefore important that we gain a better
understanding of the metabolome and proteome of the grass apoplastic space, in the
presence and absence of endophyte, to identify candidate metabolites and proteins
that are involved in signalling between the two partners. A defining feature of this
symbiotic interaction is the tight attachment of hyphae to the host cell wall (Fig. 2).
This intimate cellular connection provides an ideal passage for nutrient and signal
exchange. The fungal plasma membrane at this interface will contain a suite of
transporters and receptors that respond to various plant signals. Strategies that could
be used to refine which of these transporters and receptors are crucial for symbiotic
maintenance include RNAseq of mutant versus wild-type symbiota (Cox et al. 2010;
Eaton et al. 2010), high throughput metabolite screens using E. festucae symbiotic
gene promoters fused to EGFP (Gardiner et al. 2009), combined with ligand binding
assays, and various protein–protein interaction assays. RNAseq also has the potential to identify the suite of transcriptional regulators that are associated with the
symbiotic regulatory network. A comparison of the transcriptomes of sakA mutant
versus wild-type symbiosis identified a large set of genes encoding putative transcription factors that were differentially expressed, including Zn(II)2Cys6 binuclear
cluster, basic region-leucine zipper (bZIP) and Zn(II) coordinating Cys2His2 factors
(Eaton et al. 2010). Targeted deletion of the genes for these putative transcription
factors combined with plant phenotype testing will determine if these genes are
required for symbiotic maintenance. Genes that affect secondary metabolite biosynthesis or the symbiotic interaction with the host could be further analysed by a
combination of ChIPseq and RNAseq to identify the downstream effectors of these
transcriptional regulators. These combined approaches will provide important
insights into signalling in the E. festucae–L. perenne symbiosis and help understand
the differences between fungal mutualistic and pathogenic interactions with plants.
Acknowledgements This research was supported by grants from the Tertiary Education Commission (TEC) to the Bio-Protection Research Centre, the Royal Society of New Zealand Marsden
Fund (07MAU046), Massey University and by a Top Achiever Doctoral Scholarship to CE from
TEC. We thank Emma Brasell (IMBS) and Dimitry Sokolov (Manawatu Microscopy and Imaging
Centre, IMBS) for preparation of the image used in Fig. 1.
The E. festucae E2368 and E894 genome sequences were made available by Christopher
Schardl through grants EF-0523661 from the US National Science Foundation (to Christopher
Schardl, Mark Farman and Bruce Roe) and 2005-35319-16141 from the US Department of
Agriculture National Research (to Christopher Schardl).
References
Adachi K, Hamer JE (1998) Divergent cAMP signaling pathways regulate growth and pathogenesis in the rice blast fungus Magnaporthe grisea. Plant Cell 10:1361–1374
Alex LA, Borkovich KA, Simon MI (1996) Hyphal development in Neurospora crassa: involvement of a two-component histidine kinase. Proc Natl Acad Sci USA 93:3416–3421
Allen RG, Tresini M (2000) Oxidative stress and gene regulation. Free Radic Biol Med
28:463–499
Signalling in the Epichloe¨ festucae: Perennial Ryegrass Mutualistic Symbiotic Interaction
173
Baier M, Kandlbinder A, Golldack D, Dietz K-J (2005) Oxidative stress and ozone: perception,
signalling and response. Plant Cell Environ 28:1012–1020
Banno S, Noguchi R, Yamashita K, Fukumori F, Kimura M, Yamaguchi I, Fujimura M (2007)
Roles of putative His-to-Asp signaling modules HPT-1 and RRG-2, on viability and sensitivity
to osmotic and oxidative stresses in Neurospora crassa. Curr Genet 51:197–208
Banuett F (1998) Signaling in the yeasts: an informational cascade with links to filamentous fungi.
Microbiol Mol Biol Rev 62:249–274
Blankenship JD, Spiering MJ, Wilkinson HH, Fannin FF, Bush LP, Schardl CL (2001) Production
of loline alkaloids by the grass endophyte, Neotyphodium uncinatum, in defined media.
Phytochemistry 58:395–401
Bok JW, Chiang YM, Szewczyk E, Reyes-Dominguez Y, Davidson AD, Sanchez JF, Lo HC,
Watanabe K, Strauss J, Oakley BR, Wang CC, Keller NP (2009) Chromatin-level regulation of
biosynthetic gene clusters. Nat Chem Biol 5:462–464
Bootman MD, Collins TJ, Peppiatt CM, Prothero LS, MacKenzie L, De Smet P, Travers M, Tovey
SC, Seo JT, Berridge MJ, Ciccolini F, Lipp P (2001) Calcium signalling – an overview. Semin
Cell Dev Biol 12:3–10
Bormann J, Tudzynski P (2009) Deletion of MID1, a putative stretch-activated calcium channel in
Claviceps purpurea, affects vegetative growth, cell wall synthesis and virulence. Microbiology
155:3922–3933
Brewster JL, de Valoir T, Dwyer ND, Winter E, Gustin MC (1993) An osmosensing signal
transduction pathway in yeast. Science 259:1760–1763
Bruno KS, Aramayo R, Minke PF, Metzenberg RL, Plamann M (1996) Loss of growth polarity and
mislocalization of septa in a Neurospora mutant altered in the regulatory subunit of cAMPdependent protein kinase. EMBO J 15:5772–5782
Buchanan BB, Balmer Y (2005) Redox regulation: a broadening horizon. Annu Rev Plant Biol
56:187–220
Cano-Domı́nguez N, Álvarez-Delfı́n K, Hansberg W, Aguirre J (2008) The NADPH oxidases
NOX-1 and NOX-2 require the regulatory subunit NOR-1 to control cell differentiation and
growth in Neurospora crassa. Eukaryot Cell 7:1352–1361
Chang C, Stewart RC (1998) The two-component system. Regulation of diverse signaling
pathways in prokaryotes and eukaryotes. Plant Physiol 117:723–731
Chen SX, Schopfer P (1999) Hydroxyl-radical production in physiological reactions. A novel
function of peroxidase. Eur J Biochem 260:726–735
Cheng H, Lederer WJ (2008) Calcium sparks. Physiol Rev 88:1491–1545
Choi W, Dean RA (1997) The adenylate cyclase gene MAC1 of Magnaporthe grisea controls
appressorium formation and other aspects of growth and development. Plant Cell 9:1973–1983
Choi YE, Xu JR (2010) The cAMP signaling pathway in Fusarium verticillioides is important for
conidiation, plant infection, and stress responses but not fumonisin production. Mol Plant
Microbe Interact 23:522–533
Choi J, Kim Y, Kim S, Park J, Lee YH (2009a) MoCRZ1, a gene encoding a calcineurinresponsive transcription factor, regulates fungal growth and pathogenicity of Magnaporthe
oryzae. Fungal Genet Biol 46:243–254
Choi JH, Kim Y, Lee YH (2009b) Functional analysis of MCNA, a gene encoding a catalytic
subunit of calcineurin, in the rice blast fungus Magnaporthe oryzae. J Microbiol Biotechnol
19:11–16
Christensen MJ, Ball OJ-P, Bennett RJ, Schardl CL (1997) Fungal and host genotype effects on
compatibility and vascular colonization by Epichloe¨ festucae. Mycol Res 101:493–501
Christensen MJ, Bennett RJ, Ansari HA, Koga H, Johnson RD, Bryan GT, Simpson WR, Koolaard
JP, Nickless EM, Voisey CR (2008) Epichloe¨ endophytes grow by intercalary hyphal extension
in elongating grass leaves. Fungal Genet Biol 45:84–93
Cox MP, Eaton CJ, Scott B (2010) Exploring molecular signaling in plant-fungal symbioses using
high throughput RNA sequencing. Plant Signal Behav 5:1353–1358
Cunningham KW, Fink GR (1996) Calcineurin inhibits VCX1-dependent H+/Ca2+ exchange and
induces Ca2+ ATPases in Saccharomyces cerevisiae. Mol Cell Biol 16:2226–2237
174
C. Eaton et al.
D’Souza CA, Heitman J (2001) Conserved cAMP signaling cascades regulate fungal development
and virulence. FEMS Microbiol Rev 25:349–364
Davis TN, Urdea MS, Masiarz FR, Thorner J (1986) Isolation of the yeast calmodulin gene:
calmodulin is an essential protein. Cell 47:423–431
Dayton JS, Means AR (1996) Ca2+/calmodulin-dependent kinase is essential for both growth and
nuclear division in Aspergillus nidulans. Mol Biol Cell 7:1511–1519
Dayton JS, Sumi M, Nanthakumar NN, Means AR (1997) Expression of a constitutively active Ca2+/
calmodulin-dependent kinase in Aspergillus nidulans spores prevents germination and entry into
the cell cycle. J Biol Chem 272:3223–3230
Dean RA, Talbot NJ, Ebbole DJ, Farman ML, Mitchell TK, Orbach MJ, Thon M, Kulkarni R, Xu
JR, Pan H, Read ND, Lee YH, Carbone I, Brown D, Oh YY, Donofrio N, Jeong JS, Soanes DM,
Djonovic S, Kolomiets E, Rehmeyer C, Li W, Harding M, Kim S, Lebrun MH, Bohnert H,
Coughlan S, Butler J, Calvo S, Ma LJ, Nicol R, Purcell S, Nusbaum C, Galagan JE and Birren
BW (2005) The genome sequence of the rice blast fungus Magnaporthe grisea. Nature 434:
980–986
Delaunay A, Pflieger D, Barrault MB, Vinh J, Toledano MB (2002) A thiol peroxidase is an H2O2
receptor and redox-transducer in gene activation. Cell 111:471–481
Denis V, Cyert MS (2002) Internal Ca2+ release in yeast is triggered by hypertonic shock and
mediated by a TRP channel homologue. J Cell Biol 156:29–34
Di Pietro A, Garcia-MacEira FI, Meglecz E, Roncero MI (2001) A MAP kinase of the vascular
wilt fungus Fusarium oxysporum is essential for root penetration and pathogenesis. Mol
Microbiol 39:1140–1152
Di Pietro A, Madrid MP, Caracuel Z, Delgado-Jarana J, Roncero MI (2003) Fusarium oxysporum:
exploring the molecular arsenal of a vascular wilt fungus. Mol Plant Pathol 4:315–325
Dixon KP, Xu JR, Smirnoff N, Talbot NJ (1999) Independent signaling pathways regulate cellular
turgor during hyperosmotic stress and appressorium-mediated plant infection by Magnaporthe
grisea. Plant Cell 11:2045–2058
Durrenberger F, Wong K, Kronstad JW (1998) Identification of a cAMP-dependent protein kinase
catalytic subunit required for virulence and morphogenesis in Ustilago maydis. Proc Natl Acad
Sci USA 95:5684–5689
Dutta R, Robinson KR (2004) Identification and characterization of stretch-activated ion channels
in pollen protoplasts. Plant Physiol 135:1398–1406
Eaton CJ, Cox MP, Ambrose B, Becker M, Hesse U, Schardl CL, Scott B (2010) Disruption of
signaling in a fungal-grass symbiosis leads to pathogenesis. Plant Physiol 153:1780–1794
Egan JD, Garcia-Pedrajas MD, Andrews DL, Gold SE (2009) Calcineurin is an antagonist to PKA
protein phosphorylation required for postmating filamentation and virulence, while PP2A is
required for viability in Ustilago maydis. Mol Plant Microbe Interact 22:1293–1301
Feijo JA, Sainhas J, Holdaway-Clarke T, Cordeiro MS, Kunkel JG, Hepler PK (2001) Cellular
oscillations and the regulation of growth: the pollen tube paradigm. Bioessays 23:86–94
Fleetwood DJ, Scott B, Lane GA, Tanaka A, Johnson RD (2007) A complex ergovaline gene
cluster in epichloë endophytes of grasses. Appl Environ Microbiol 73:2571–2579
Flick JS, Thorner J (1993) Genetic and biochemical characterization of a phosphatidylinositolspecific phospholipase C in Saccharomyces cerevisiae. Mol Cell Biol 13:5861–5876
Foreman J, Demidchik V, Bothwell JH, Mylona P, Miedema H, Torres MA, Linstead P, Costa S,
Brownlee C, Jones JD, Davies JM, Dolan L (2003) Reactive oxygen species produced by
NADPH oxidase regulate plant cell growth. Nature 422:442–446
Forman HJ, Fukuto JM, Torres M (2004) Redox signaling: thiol chemistry defines which reactive
oxygen and nitrogen species can act as second messengers. Am J Physiol Cell Physiol
287:246–256
Galagan JE, Calvo SE, Borkovich KA, Selker EU, Read ND, Jaffe D, FitzHugh W, Ma LJ,
Smirnov S, Purcell S, Rehman B, Elkins T, Engels R, Wang S, Nielsen CB, Butler J,
Endrizzi M, Qui D, Ianakiev P, Bell-Pedersen D, Nelson MA, Werner-Washburne M,
Selitrennikoff CP, Kinsey JA, Braun EL, Zelter A, Schulte U, Kothe GO, Jedd G, Mewes W,
Staben C, Marcotte E, Greenberg D, Roy A, Foley K, Naylor J, Stange-Thomann N, Barrett R,
Signalling in the Epichloe¨ festucae: Perennial Ryegrass Mutualistic Symbiotic Interaction
175
Gnerre S, Kamal M, Kamvysselis M, Mauceli E, Bielke C, Rudd S, Frishman D, Krystofova S,
Rasmussen C, Metzenberg RL, Perkins DD, Kroken S, Cogoni C, Macino G, Catcheside D, Li
W, Pratt RJ, Osmani SA, DeSouza CP, Glass L, Orbach MJ, Berglund JA, Voelker R,
Yarden O, Plamann M, Seiler S, Dunlap J, Radford A, Aramayo R, Natvig DO, Alex LA,
Mannhaupt G, Ebbole DJ, Freitag M, Paulsen I, Sachs MS, Lander ES, Nusbaum C, Birren B
(2003) The genome sequence of the filamentous fungus Neurospora crassa. Nature
422:859–868
Ganem S, Lu SW, Lee BN, Chou DY, Hadar R, Turgeon BG, Horwitz BA (2004) G-protein b
subunit of Cochliobolus heterostrophus involved in virulence, asexual and sexual reproductive
ability, and morphogenesis. Eukaryot Cell 3:1653–1663
Gao S, Nuss DL (1996) Distinct roles for two G protein a subunits in fungal virulence, morphology, and reproduction revealed by targeted gene disruption. Proc Natl Acad Sci USA
93:14122–14127
Gardiner DM, Kazan K, Manners JM (2009) Nutrient profiling reveals potent inducers of trichothecene biosynthesis in Fusarium graminearum. Fungal Genet Biol 46:604–613
Gold S, Duncan G, Barrett K, Kronstad J (1994) cAMP regulates morphogenesis in the fungal
pathogen Ustilago maydis. Genes Dev 8:2805–2816
Grissa I, Bidard F, Grognet P, Grossetete S, Silar P (2010) The Nox/Ferric reductase/Ferric
reductase-like families of Eumycetes. Fungal Biol 114:766–777
Groemping Y, Rittinger K (2005) Activation and assembly of the NADPH oxidase: a structural
perspective. Biochem J 386:401–416
Gronover CS, Kasulke D, Tudzynski P, Tudzynski B (2001) The role of G protein alpha subunits
in the infection process of the gray mold fungus Botrytis cinerea. Mol Plant Microbe Interact
14:1293–1302
Gustin MC, Albertyn J, Alexander M, Davenport K (1998) MAP kinase pathways in the yeast
Saccharomyces cerevisiae. Microbiol Mol Biol Rev 62:1264–1300
Halachmi D, Eilam Y (1989) Cytosolic and vacuolar Ca2+ concentrations in yeast cells measured
with the Ca2+-sensitive fluorescence dye indo-1. FEBS Lett 256:55–61
Hancock JT, Desikan R, Neill SJ (2001) Role of reactive oxygen species in cell signalling
pathways. Biochem Soc Trans 29:345–350
Heller G, Adomas A, Li G, Osborne J, van Zyl L, Sederoff R, Finlay RD, Stenlid J, Asiegbu FO
(2008) Transcriptional analysis of Pinus sylvestris roots challenged with the ectomycorrhizal
fungus Laccaria bicolor. BMC Plant Biol 8:19
Horwitz BA, Sharon A, Lu SW, Ritter V, Sandrock TM, Yoder OC, Turgeon BG (1999) A G
protein alpha subunit from Cochliobolus heterostrophus involved in mating and appressorium
formation. Fungal Genet Biol 26:19–32
Hou Z, Xue C, Peng Y, Katan T, Kistler HC, Xu JR (2002) A mitogen-activated protein kinase
gene (MGV1) in Fusarium graminearum is required for female fertility, heterokaryon formation, and plant infection. Mol Plant Microbe Interact 15:1119–1127
Igbaria A, Lev S, Rose MS, Lee BN, Hadar R, Degani O, Horwitz BA (2008) Distinct and
combined roles of the MAP kinases of Cochliobolus heterostrophus in virulence and stress
responses. Mol Plant Microbe Interact 21:769–780
Iida H, Nakamura H, Ono T, Okumura MS, Anraku Y (1994) MID1, a novel Saccharomyces
cerevisiae gene encoding a plasma membrane protein, is required for Ca2+ influx and mating.
Mol Cell Biol 14:8259–8271
Iida H, Ohya Y, Anraku Y (1995) Calmodulin-dependent protein kinase II and calmodulin are
required for induced thermotolerance in Saccharomyces cerevisiae. Curr Genet 27:190–193
Jackson SL, Heath IB (1993) Roles of calcium ions in hyphal tip growth. Microbiol Rev
57:367–382
Jagnandan D, Church JE, Banfi B, Stuehr DJ, Marrero MB, Fulton DJ (2007) Novel mechanism of
activation of NADPH oxidase 5. Calcium sensitization via phosphorylation. J Biol Chem
282:6494–6507
176
C. Eaton et al.
Jain S, Akiyama K, Mae K, Ohguchi T, Takata R (2002) Targeted disruption of a G protein a
subunit gene results in reduced pathogenicity in Fusarium oxysporum. Curr Genet 41:407–413
Jain S, Akiyama K, Kan T, Ohguchi T, Takata R (2003) The G protein b subunit FGB1 regulates
development and pathogenicity in Fusarium oxysporum. Curr Genet 43:79–86
Jeon J, Goh J, Yoo S, Chi MH, Choi J, Rho HS, Park J, Han SS, Kim BR, Park SY, Kim S, Lee YH
(2008) A putative MAP kinase kinase kinase, MCK1, is required for cell wall integrity and
pathogenicity of the rice blast fungus, Magnaporthe oryzae. Mol Plant Microbe Interact
21:525–534
Johnson L (2008) Iron and siderophores in fungal-host interactions. Mycol Res 112:170–183
Johnson LJ, Steringa M, Koulman A, Christensen M, Johnson RD, Voisey CR, Bryan G, Lamont I,
Rasmussen S (2007) Biosynthesis of an extracellular siderophore is essential for maintenance
of mutualistic endophyte-grass symbioses. In: Popay AJ, Thon ER (eds) Proceedings of the
sixth international symposium on fungal endophytes of grasses grassland research and practice
series no 13. New Zealand Grassland Association, Christchurch, New Zealand, pp 177–179
Jones JD, Dangl JL (2006) The plant immune system. Nature 444:323–329
Jones CA, Greer-Phillips SE, Borkovich KA (2007) The response regulator RRG-1 functions
upstream of a mitogen-activated protein kinase pathway impacting asexual development,
female fertility, osmotic stress, and fungicide resistance in Neurospora crassa. Mol Biol Cell
18:2123–2136
Joseph JD, Means AR (2000) Identification and characterization of two Ca2+/CaM-dependent
protein kinases required for normal nuclear division in Aspergillus nidulans. J Biol Chem
275:38230–38238
Kahmann R, Basse C, Feldbrugge M (1999) Fungal-plant signalling in the Ustilago maydis-maize
pathosystem. Curr Opin Microbiol 2:647–650
Kaku H, Nishizawa Y, Ishii-Minami N, Akimoto-Tomiyama C, Dohmae N, Takio K, Minami E,
Shibuya N (2006) Plant cells recognize chitin fragments for defense signaling through a plasma
membrane receptor. Proc Natl Acad Sci USA 103:11086–11091
Kasahara S, Nuss DL (1997) Targeted disruption of a fungal G-protein b subunit gene results in
increased vegetative growth but reduced virulence. Mol Plant Microbe Interact 10:984–993
Kawano CY, Said S (2005) Hyperbranching induced by cold-shock or snow-flake mutation in
Neurospora crassa is prevented by addition of exogenous calcium. J Basic Microbiol
45:199–206
Kim YK, Li D, Kolattukudy PE (1998) Induction of Ca2+-calmodulin signaling by hard-surface
contact primes Colletotrichum gloeosporioides conidia to germinate and form appressoria.
J Bacteriol 180:5144–5150
Kim SO, Merchant K, Nudelman R, Beyer WF Jr, Keng T, DeAngelo J, Hausladen A, Stamler JS
(2002) OxyR: a molecular code for redox-related signaling. Cell 109:383–396
Klimpel A, Gronover CS, Williamson B, Stewart JA, Tudzynski B (2002) The adenylate cyclase
(BAC) in Botrytis cinerea is required for full pathogenicity. Mol Plant Pathol 3:439–450
Kojima K, Kikuchi T, Takano Y, Oshiro E, Okuno T (2002) The mitogen-activated protein kinase
gene MAF1 is essential for the early differentiation phase of appressorium formation in
Colletotrichum lagenarium. Mol Plant Microbe Interact 15:1268–1276
Kojima K, Takano Y, Yoshimi A, Tanaka C, Kikuchi T, Okuno T (2004) Fungicide activity
through activation of a fungal signalling pathway. Mol Microbiol 53:1785–1796
Lalucque H, Silar P (2003) NADPH oxidase: an enzyme for multicellularity? Trends Microbiol
11:9–12
Lambeth JD (2004) NOX enzymes and the biology of reactive oxygen. Nat Rev Immunol
4:181–189
Lanver D, Mendoza-Mendoza A, Brachmann A, Kahmann R (2010) Sho1 and Msb2-related
proteins regulate appressorium development in the smut fungus Ustilago maydis. Plant Cell
22:2085–2101
Lardy B, Bof M, Aubry L, Paclet MH, Morel F, Satre M, Klein G (2005) NADPH oxidase
homologs are required for normal cell differentiation and morphogenesis in Dictyostelium
discoideum. Biochim Biophys Acta 1744:199–212
Signalling in the Epichloe¨ festucae: Perennial Ryegrass Mutualistic Symbiotic Interaction
177
Leslie NR, Lindsay Y, Ross SH, Downes CP (2004) Redox regulation of phosphatase function.
Biochem Soc Trans 32:1018–1020
Leuchtmann A, Schardl CL, Siegel MR (1994) Sexual compatibility and taxonomy of a new
species of Epichloe¨ symbiotic with fine fescue grasses. Mycologia 86:802–812
Lev S, Sharon A, Hadar R, Ma H, Horwitz BA (1999) A mitogen-activated protein kinase of the
corn leaf pathogen Cochliobolus heterostrophus is involved in conidiation, appressorium
formation, and pathogenicity: diverse roles for mitogen-activated protein kinase homologs in
foliar pathogens. Proc Natl Acad Sci USA 96:13542–13547
Levin DE (2005) Cell wall integrity signaling in Saccharomyces cerevisiae. Microbiol Mol Biol
Rev 69:262–291
Li L, Wright SJ, Krystofova S, Park G, Borkovich KA (2007) Heterotrimeric G protein signaling
in filamentous fungi. Annu Rev Microbiol 61:423–452
Lin CH, Yang SL, Wang NY, Chung KR (2010) The FUS3 MAPK signaling pathway of the citrus
pathogen Alternaria alternata functions independently or cooperatively with the fungal redoxresponsive AP1 regulator for diverse developmental, physiological and pathogenic processes.
Fungal Genet Biol 47:381–391
Liu S, Dean RA (1997) G protein a subunit genes control growth, development, and pathogenicity
of Magnaporthe grisea. Mol Plant Microbe Interact 10:1075–1086
Liu ZM, Kolattukudy PE (1999) Early expression of the calmodulin gene, which precedes
appressorium formation in Magnaporthe grisea, is inhibited by self-inhibitors and requires
surface attachment. J Bacteriol 181:3571–3577
Liu XH, Lu JP, Dong B, Gu Y, Lin FC (2009) Disruption of MoCMK1, encoding a putative
calcium/calmodulin-dependent kinase, in Magnaporthe oryzae. Microbiol Res 165:402–410
Ma ZB, Zhao JX, Wang LA, Zheng XB (2009) Cloning, prokaryotic expression, and bioactivity of
the calmodulin gene of Magnaporthe grisea. FEMS Microbiol Lett 300:107–114
Martienssen RA, Colot V (2001) DNA methylation and epigenetic inheritance in plants and
filamentous fungi. Science 293:1070–1074
May KJ, Bryant MK, Zhang X, Ambrose B, Scott B (2008) Patterns of expression of a lolitrem
biosynthetic gene in the Epichloe¨ festucae-perennial ryegrass symbiosis. Mol Plant Microbe
Interact 21:188–197
Mehrabi R, Kema GH (2006) Protein kinase A subunits of the ascomycete pathogen
Mycosphaerella graminicola regulate asexual fructification, filamentation, melanization and
osmosensing. Mol Plant Pathol 7:565–577
Mehrabi R, Zwiers LH, de Waard MA, Kema GH (2006) MgHog1 regulates dimorphism and
pathogenicity in the fungal wheat pathogen Mycosphaerella graminicola. Mol Plant Microbe
Interact 19:1262–1269
Mehrabi R, Ben M’Barek S, van der Lee TA, Waalwijk C, de Wit PJ, Kema GH (2009) Ga and Gb
proteins regulate the cyclic AMP pathway that is required for development and pathogenicity
of the phytopathogen Mycosphaerella graminicola. Eukaryot Cell 8:1001–1013
Mey G, Oeser B, Lebrun MH, Tudzynski P (2002) The biotrophic, non-appressorium-forming
grass pathogen Claviceps purpurea needs a Fus3/Pmk1 homologous mitogen-activated protein
kinase for colonization of rye ovarian tissue. Mol Plant Microbe Interact 15:303–312
Millar JB, Buck V, Wilkinson MG (1995) Pyp1 and Pyp2 PTPases dephosphorylate an
osmosensing MAP kinase controlling cell size at division in fission yeast. Genes Dev
9:2117–2130
Mitchell TK, Dean RA (1995) The cAMP-dependent protein kinase catalytic subunit is required
for appressorium formation and pathogenesis by the rice blast pathogen Magnaporthe grisea.
Plant Cell 7:1869–1978
Mitic M (2011) Investigation of the molecular basis of symbiosis between Epichloe festucae and
perennial ryegrass. Massey University, Palmerston North, New Zealand
Moon CD, Craven KD, Leuchtmann A, Clement SL, Schardl CL (2004) Prevalence of interspecific hybrids amongst asexual fungal endophytes of grasses. Mol Ecol 13:1455–1467
178
C. Eaton et al.
Mori IC, Schroeder JI (2004) Reactive oxygen species activation of plant Ca2+ channels. A
signaling mechanism in polar growth, hormone transduction, stress signaling, and hypothetically mechanotransduction. Plant Physiol 135:702–708
Moriwaki A, Kubo E, Arase S, Kihara J (2006) Disruption of SRM1, a mitogen-activated protein
kinase gene, affects sensitivity to osmotic and ultraviolet stressors in the phytopathogenic
fungus Bipolaris oryzae. FEMS Microbiol Lett 257:253–261
Moser MJ, Geiser JR, Davis TN (1996) Ca2+-calmodulin promotes survival of pheromone-induced
growth arrest by activation of calcineurin and Ca2+-calmodulin-dependent protein kinase. Mol
Cell Biol 16:4824–4831
Motoyama T, Kadokura K, Ohira T, Ichiishi A, Fujimura M, Yamaguchi I, Kudo T (2005) A twocomponent histidine kinase of the rice blast fungus is involved in osmotic stress response and
fungicide action. Fungal Genet Biol 42:200–212
Motoyama T, Ochiai N, Morita M, Iida Y, Usami R, Kudo T (2008) Involvement of putative
response regulator genes of the rice blast fungus Magnaporthe oryzae in osmotic stress
response, fungicide action, and pathogenicity. Curr Genet 54:185–195
Nathues E, J€orgens C, Lorenz N, Tudzynski P (2007) The histidine kinase CpHK2 has impact on
spore germination, oxidative stress and fungicide resistance, and virulence of the ergot fungus
Claviceps purpurea. Mol Plant Pathol 8:653–665
Neill SJ, Desikan R, Clarke A, Hurst RD, Hancock JT (2002) Hydrogen peroxide and nitric oxide
as signalling molecules in plants. J Exp Bot 53:1237–1247
Nguyen QB, Kadotani N, Kasahara S, Tosa Y, Mayama S, Nakayashiki H (2008) Systematic
functional analysis of calcium-signalling proteins in the genome of the rice-blast fungus,
Magnaporthe oryzae, using a high-throughput RNA-silencing system. Mol Microbiol
68:1348–1365
Nishimura M, Park G, Xu JR (2003) The G-beta subunit MGB1 is involved in regulating multiple
steps of infection-related morphogenesis in Magnaporthe grisea. Mol Microbiol 50:231–243
Ohya Y, Kawasaki H, Suzuki K, Londesborough J, Anraku Y (1991) Two yeast genes encoding
calmodulin-dependent protein kinases. Isolation, sequencing and bacterial expression of
CMK1 and CMK2. J Biol Chem 266:12784–12794
Oide S, Liu J, Yun SH, Wu D, Michev A, Choi MY, Horwitz BA, Turgeon BG (2010) Histidine
kinase two component response regulator proteins regulate reproductive development, virulence, and stress responses of the fungal cereal pathogens Cochliobolus heterostrophus and
Gibberella zeae. Eukaryot Cell 9:1867–1880
Paidhungat M, Garrett S (1997) A homolog of mammalian, voltage-gated calcium channels
mediates yeast pheromone-stimulated Ca2+ uptake and exacerbates the cdc1(Ts) growth defect.
Mol Cell Biol 17:6339–6347
Palmer CP, Zhou XL, Lin J, Loukin SH, Kung C, Saimi Y (2001) A TRP homolog in Saccharomyces cerevisiae forms an intracellular Ca2+-permeable channel in the yeast vacuolar membrane. Proc Natl Acad Sci USA 98:7801–7805
Panaccione DG, Johnson RD, Wang J, Young CA, Damrongkool P, Scott B, Schardl CL (2001)
Elimination of ergovaline from a grass-Neotyphodium endophyte symbiosis by genetic modification of the endophyte. Proc Natl Acad Sci USA 98:12820–12825
Park SY, Seo SB, Lee SJ, Na JG, Kim YJ (2001) Mutation in PMR1, a Ca2+-ATPase in Golgi,
confers salt tolerance in Saccharomyces cerevisiae by inducing expression of PMR2, an Na+ATPase in plasma membrane. J Biol Chem 276:28694–28699
Pei ZM, Murata Y, Benning G, Thomine S, Klusener B, Allen GJ, Grill E, Schroeder JI (2000)
Calcium channels activated by hydrogen peroxide mediate abscisic acid signalling in guard
cells. Nature 406:731–734
Permyakov EA, Kretsinger RH (2009) Cell signaling, beyond cytosolic calcium in eukaryotes.
J Inorg Biochem 103:77–86
Philipson MN, Christey MC (1986) The relationship of host and endophyte during flowering, seed
formation, and germination of Lolium perenne. NZ J Bot 24:125–134
Pinchai N, Juvvadi PR, Fortwendel JR, Perfect BZ, Rogg LE, Asfaw YG, Steinbach WJ (2010)
The Aspergillus fumigatus P-type Golgi apparatus Ca2+/Mn2+ ATPase PmrA is involved in
Signalling in the Epichloe¨ festucae: Perennial Ryegrass Mutualistic Symbiotic Interaction
179
cation homeostasis and cell wall integrity but is not essential for pathogenesis. Eukaryot Cell
9:472–476
Prokisch H, Yarden O, Dieminger M, Tropschug M, Barthelmess IB (1997) Impairment of
calcineurin function in Neurospora crassa reveals its essential role in hyphal growth, morphology and maintenance of the apical Ca2+ gradient. Mol Gen Genet 256:104–114
Regenfelder E, Spellig T, Hartmann A, Lauenstein S, Bolker M, Kahmann R (1997) G proteins in
Ustilago maydis: transmission of multiple signals? EMBO J 16:1934–1942
Reyes-Dominguez Y, Bok JW, Berger H, Shwab EK, Basheer A, Gallmetzer A, Scazzocchio C,
Keller N, Strauss J (2010) Heterochromatic marks are associated with the repression of
secondary metabolism clusters in Aspergillus nidulans. Mol Microbiol 76:1376–1386
Rho HS, Jeon J, Lee YH (2009) Phospholipase C-mediated calcium signalling is required for
fungal development and pathogenicity in Magnaporthe oryzae. Mol Plant Pathol 10:337–346
Rispail N, Di Pietro A (2010) The two-component histidine kinase Fhk1 controls stress adaptation
and virulence of Fusarium oxysporum. Mol Plant Pathol 11:395–407
Rispail N, Soanes DM, Ant C, Czajkowski R, Grunler A, Huguet R, Perez-Nadales E, Poli A,
Sartorel E, Valiante V, Yang M, Beffa R, Brakhage AA, Gow NA, Kahmann R, Lebrun MH,
Lenasi H, Perez-Martin J, Talbot NJ, Wendland J, Di Pietro A (2009) Comparative genomics
of MAP kinase and calcium-calcineurin signalling components in plant and human pathogenic
fungi. Fungal Genet Biol 46:287–298
Rolke Y, Tudzynski P (2008) The small GTPase Rac and the p21-activated kinase Cla4 in
Claviceps purpurea: interaction and impact on polarity, development and pathogenicity. Mol
Microbiol 68:405–423
Rusnak F, Mertz P (2000) Calcineurin: form and function. Physiol Rev 80:1483–1521
Saito H, Tatebayashi K (2004) Regulation of the osmoregulatory HOG MAPK cascade in yeast.
J Biochem 136:267–272
Saudohar M, Bencina M, van de Vondervoort PJ, Panneman H, Legisa M, Visser J, Ruijter GJ
(2002) Cyclic AMP-dependent protein kinase is involved in morphogenesis of Aspergillus
niger. Microbiology 148:2635–2645
Schardl CL, Wilkinson HH (2000) Hybridization and cospeciation hypotheses for the evolution of
grass endophytes. In: Bacon CW, White JF Jr (eds) Microbial endophytes. Marcel Dekker, Inc.,
New York, pp 63–83
Schardl CL, Panaccione DG, Tudzynski P (2006) Ergot alkaloids-biology and molecular biology.
In: Cordell GA (ed) Alkaloids: chemistry and biology. Academic, San Diego, CA, pp 425–443
Schmid J, Harold FM (1988) Dual roles for calcium ions in apical growth of Neurospora crassa.
J Gen Microbiol 134:2623–2631
Schumacher J, de Larrinoa IF, Tudzynski B (2008) Calcineurin-responsive zinc finger transcription factor CRZ1 of Botrytis cinerea is required for growth, development, and full virulence on
bean plants. Eukaryot Cell 7:584–601
Scott B, Takemoto D, Tanaka A, Young CA, Bryant MK, May KJ (2007) Functional analysis of the
Epichloë festucae-perennial ryegrass symbiosis. In: Popay AJ, Thom ER (eds) Proceedings of the
sixth international symposium on fungal endophytes of grasses grasslands research and practice,
series no 13. New Zealand Grassland Association, Christchurch, New Zealand, pp 433–441
Segm€uller N, Ellendorf U, Tudzynski B, Tudzynski P (2007) BcSAK1, a stress-activated mitogenactivated protein kinase, is involved in vegetative differentiation and pathogenicity in Botrytis
cinerea. Eukaryot Cell 6:211–221
Semighini CP, Harris SD (2008) Regulation of apical dominance in Aspergillus nidulans hyphae
by reactive oxygen species. Genetics 179:1919–1932
Shiozaki K, Russell P (1995) Cell-cycle control linked to extracellular environment by MAP
kinase pathway in fission yeast. Nature 378:739–743
Shwab EK, Bok JW, Tribus M, Galehr J, Graessle S, Keller NP (2007) Histone deacetylase activity
regulates chemical diversity in Aspergillus. Eukaryot Cell 6:1656–1664
Silverman-Gavrila LB, Lew RR (2002) An IP3-activated Ca2+ channel regulates fungal tip growth.
J Cell Sci 115:5013–5025
180
C. Eaton et al.
Silverman-Gavrila LB, Lew RR (2003) Calcium gradient dependence of Neurospora crassa
hyphal growth. Microbiology 149:2475–2485
Smith DG, Garcia-Pedrajas MD, Hong W, Yu Z, Gold SE, Perlin MH (2004) An ste20 homologue
in Ustilago maydis plays a role in mating and pathogenicity. Eukaryot Cell 3:180–189
Solomon PS, Tan KC, Sanchez P, Cooper RM, Oliver RP (2004) The disruption of a Ga subunit
sheds new light on the pathogenicity of Stagonospora nodorum on wheat. Mol Plant Microbe
Interact 17:456–466
Solomon PS, Rybak K, Trengove RD, Oliver RP (2006) Investigating the role of calcium/
calmodulin-dependent protein kinases in Stagonospora nodorum. Mol Microbiol 62:367–381
Spiering MJ, Moon CD, Wilkinson HH, Schardl CL (2005) Gene clusters for insecticidal loline
alkaloids in the grass-endophytic fungus Neotyphodium uncinatum. Genetics 169:1403–1414
Stie J, Fox D (2008) Calcineurin regulation in fungi and beyond. Eukaryot Cell 7:177–186
Stock AM, Robinson VL, Goudreau PN (2000) Two-component signal transduction. Annu Rev
Biochem 69:183–215
Sumimoto H, Miyano K, Takeya R (2005) Molecular composition and regulation of the Nox
family NAD(P)H oxidases. Biochem Biophys Res Commun 338:677–686
Taglialatela M, Castaldo P, Iossa S, Pannaccione A, Fresi A, Ficker E, Annunziato L (1997)
Regulation of the human ether-a-gogo related gene (HERG) K+ channels by reactive oxygen
species. Proc Natl Acad Sci USA 94:11698–11703
Takano Y, Kikuchi T, Kubo Y, Hamer JE, Mise K, Furusawa I (2000) The Colletotrichum
lagenarium MAP kinase gene CMK1 regulates diverse aspects of fungal pathogenesis. Mol
Plant Microbe Interact 13:374–383
Takeda K, Iizuka M, Watanabe T, Nakagawa J, Kawasaki S, Niimura Y (2007) Synechocystis
DrgA protein functioning as nitroreductase and ferric reductase is capable of catalyzing the
Fenton reaction. FEBS J 274:1318–1327
Takeda S, Gapper C, Kaya H, Bell E, Kuchitsu K, Dolan L (2008) Local positive feedback
regulation determines cell shape in root hair cells. Science 319:1241–1244
Takemoto D, Tanaka A, Scott B (2006) A p67Phox-like regulator is recruited to control hyphal
branching in a fungal-grass mutualistic symbiosis. Plant Cell 18:2807–2821
Takemoto D, Tanaka A, Scott B (2007) NADPH oxidases in fungi: diverse roles of reactive
oxygen species in fungal cellular differentiation. Fungal Genet Biol 44:1065–1076
Takemoto D, Kamakura S, Saikia S, Becker Y, Wrenn R, Tanaka A, Sumimoto H, Scott B (2011)
Polarity proteins Bem1 and Cdc24 are components of the filamentous fungal NADPH oxidase
complex. Proc Natl Acad Sci USA 108:2861–2866
Tanaka A, Tapper BA, Popay A, Parker EJ, Scott B (2005) A symbiosis expressed non-ribosomal
peptide synthetase from a mutualistic fungal endophyte of perennial ryegrass confers protection to the symbiotum from insect herbivory. Mol Microbiol 57:1036–1050
Tanaka A, Christensen MJ, Takemoto D, Park P, Scott B (2006) Reactive oxygen species play a
role in regulating a fungus-perennial ryegrass mutualistic association. Plant Cell 18:1052–1066
Tanaka A, Takemoto D, Hyon GS, Park P, Scott B (2008) NoxA activation by the small GTPase
RacA is required to maintain a mutualistic symbiotic association between Epichloe¨ festucae
and perennial ryegrass. Mol Microbiol 68:1165–1178
Tisi R, Baldassa S, Belotti F, Martegani E (2002) Phospholipase C is required for glucose-induced
calcium influx in budding yeast. FEBS Lett 520:133–138
Torralba S, Heath IB (2001) Cytoskeletal and Ca2+ regulation of hyphal tip growth and initiation.
Curr Top Dev Biol 51:135–187
Torres MA, Onouchi H, Hamada S, Machida C, Hammond-Kosack KE, Jones JDG (1998) Six
Arabidopsis thaliana homologues of the human respiratory burst oxidase (gp91phox). Plant
J 14:365–370
Turner GE, Borkovich KA (1993) Identification of a G protein alpha subunit from Neurospora
crassa that is a member of the Gi family. J Biol Chem 268:14805–14811
Vargas-Perez I, Sanchez O, Kawasaki L, Georgellis D, Aguirre J (2007) Response regulators SrrA
and SskA are central components of a phosphorelay system involved in stress signal transduction and asexual sporulation in Aspergillus nidulans. Eukaryot Cell 6:1570–1583
Signalling in the Epichloe¨ festucae: Perennial Ryegrass Mutualistic Symbiotic Interaction
181
Viaud M, Fillinger S, Liu W, Polepalli JS, Le Pecheur P, Kunduru AR, Leroux P, Legendre L
(2006) A class III histidine kinase acts as a novel virulence factor in Botrytis cinerea. Mol Plant
Microbe Interact 19:1042–1050
Voisey CR, Christensen MJ, Johnson RD, Johnson LJ, Khan AK, Park ZA, Ramakrishna M, Cao M,
Koolaard JP, Koulman A, Borchert S, Gaborit CN, Pratt JM, Rasmussen S, Bryan GT (2007)
The role of cAMP signalling in the symbiosis between Epichloe¨ festucae and Lolium perenne.
In: Popay AJ, Thom ER (eds) Proceedings of the sixth international symposium on fungal
endophytes of grasses grasslands research and practice. Series no 13. New Zealand Grassland
Association, Christchurch, New Zealand, pp 457–459
Wang J, Machado C, Panaccione DG, Tsai H-F, Schardl CL (2004) The determinant step in ergot
alkaloid biosynthesis by an endophyte of perennial ryegrass. Fungal Genet Biol 41:189–198
Xu JR, Hamer JE (1996) MAP kinase and cAMP signaling regulate infection structure formation
and pathogenic growth in the rice blast fungus Magnaporthe grisea. Genes Dev 10:2696–2706
Xu JR, Staiger CJ, Hamer JE (1998) Inactivation of the mitogen-activated protein kinase Mps1
from the rice blast fungus prevents penetration of host cells but allows activation of plant
defense responses. Proc Natl Acad Sci USA 95:12713–12718
Yan Y, Wei CL, Zhang WR, Cheng HP, Liu J (2006) Cross-talk between calcium and reactive
oxygen species signaling. Acta Pharmacol Sin 27:821–826
Yang Y, Cheng P, Zhi G, Liu Y (2001) Identification of a calcium/calmodulin-dependent protein
kinase that phosphorylates the Neurospora circadian clock protein FREQUENCY. J Biol Chem
276:41064–41072
Yoko-o T, Matsui Y, Yagisawa H, Nojima H, Uno I, Toh-e A (1993) The putative phosphoinositide-specific phospholipase C gene, PLC1, of the yeast Saccharomyces cerevisiae is
important for cell growth. Proc Natl Acad Sci USA 90:1804–1808
Young CA, Bryant MK, Christensen MJ, Tapper BA, Bryan GT, Scott B (2005) Molecular cloning
and genetic analysis of a symbiosis-expressed gene cluster for lolitrem biosynthesis from a
mutualistic endophyte of perennial ryegrass. Mol Genet Genomics 274:13–29
Young CA, Felitti S, Shields K, Spangenberg G, Johnson RD, Bryan GT, Saikia S, Scott B (2006)
A complex gene cluster for indole-diterpene biosynthesis in the grass endophyte Neotyphodium
lolii. Fungal Genet Biol 43:679–693
Zelter A, Bencina M, Bowman BJ, Yarden O, Read ND (2004) A comparative genomic analysis of
the calcium signaling machinery in Neurospora crassa, Magnaporthe grisea, and Saccharomyces cerevisiae. Fungal Genet Biol 41:827–841
Zhang H, Zhao Q, Liu K, Zhang Z, Wang Y, Zheng X (2009) MgCRZ1, a transcription factor of
Magnaporthe grisea, controls growth, development and is involved in full virulence. FEMS
Microbiol Lett 293:160–169
Zhao X, Kim Y, Park G, Xu JR (2005) A mitogen-activated protein kinase cascade regulating
infection-related morphogenesis in Magnaporthe grisea. Plant Cell 17:1317–1329
Zheng L, Campbell M, Murphy J, Lam S, Xu JR (2000) The BMP1 gene is essential for
pathogenicity in the gray mold fungus Botrytis cinerea. Mol Plant Microbe Interact
13:724–732
Zheng W, Zhao Z, Chen J, Liu W, Ke H, Zhou J, Lu G, Darvill AG, Albersheim P, Wu S, Wang Z
(2009) A Cdc42 ortholog is required for penetration and virulence of Magnaporthe grisea.
Fungal Genet Biol 46:450–460
Zhou XL, Batiza AF, Loukin SH, Palmer CP, Kung C, Saimi Y (2003) The transient receptor
potential channel on the yeast vacuole is mechanosensitive. Proc Natl Acad Sci USA
100:7105–7110
Zipfel C (2008) Pattern-recognition receptors in plant innate immunity. Curr Opin Immunol
20:10–16
Plant Infection by Biotrophic Fungal
and Oomycete Pathogens
Pamela H.P. Gan, Peter N. Dodds, and Adrienne R. Hardham
Abstract Biotrophic fungi and oomycetes constitute some of the most destructive
pathogens of agriculturally important plants. In susceptible hosts, these pathogens
infect and establish a dynamic relationship with living plant cells through which
they redirect plant resources to support pathogen growth and reproduction. During
evolution, biotrophic infection strategies have become finely tuned to allow the
pathogen to avoid or suppress the plant’s defence response and to take control of
host cell organisation and metabolism. Recent research has begun to uncover
intriguing details of pathogen effector proteins that are instrumental in facilitating
biotrophic growth. These effectors function in both the plant apoplast and cytoplasm. They enable plant penetration, they mask factors that normally trigger basal
defence and they suppress the host defence response. Effectors responsible for
redirecting host metabolism remain to be characterised. Ongoing research efforts
focus on elucidation of effector identity and function and of the mechanisms that
operate in the uptake of intracellular effectors into the host cytoplasm.
1 Introduction
Plants constantly come into contact with a diverse range of micro-organisms, be
they viruses, bacteria, fungi or oomycetes. Nevertheless, in the majority of cases
they are able to fend off infection by pathogens, making disease the exception
rather than the rule. In the case of fungi, fewer than 2% of the 100,000 fungal
P.H.P. Gan • A.R. Hardham (*)
Plant Science Center, RIKEN, Yokohama, Kanagawa 230-0045 Japan
e-mail: [email protected]; [email protected]
P.N. Dodds
Division of Plant Industry, CSIRO, Canberra, ACT 2601 Australia
e-mail: [email protected]
S. Perotto and F. Baluska (eds.), Signaling and Communication in Plant Symbiosis,
Signaling and Communication in Plants 11, DOI 10.1007/978-3-642-20966-6_8,
# Springer-Verlag Berlin Heidelberg 2012
183
184
P.H.P. Gan et al.
species known to exist actually cause disease in living plants (Agrios 1997). Of the
rest, some are mutualistic, as described in other chapters of this text, but most are
saprophytes that are strictly involved in the decomposition of plant material that is
already dead. Furthermore, those micro-organisms that are pathogenic can infect
only a limited number of host plant species (their host range). There is an on-going
evolutionary battle, with individual plants being susceptible to certain pathogenic
micro-organisms and individual pathogens able to infect only certain plant species
or cultivars. Thus, while resistance to microbial attack has evolved in most plants,
the same evolutionary forces have given rise to pathogens capable of overcoming
plant defences, often with devastating results. This chapter presents our current
understanding of the strategies fungal and oomycete pathogens use to infect plants
and establish disease.
Fungi and oomycetes are two major groups of eukaryotic plant pathogens. They
share many morphological similarities that are likely to have arisen through convergent evolution as efficient mechanisms for development and reproduction,
including growth as filamentous hyphae and production and dispersal of sexual
and asexual spores (Latijnhouwers et al. 2003). Early plant pathologists considered
oomycetes to be a class of organisms within the fungal kingdom, however, structural and biochemical differences and comparisons of nucleotide sequences have
provided strong evidence that the two groups are phylogenetically separate
(Hardham 2005; Van de Peer and De Watcher 1997). It is now recognised that
the oomycetes are more closely related to the coloured algae and alveolates, the
latter including apicomplexan parasites such as Plasmodium falciparum, the causal
agent of malaria (Baldauf et al. 2000), than they are to true fungi.
Fungal and oomycete plant pathogens have two main life-styles. Necrotrophic
pathogens infect living plants but quickly kill plant tissues, obtaining nutrients from
dead or dying cells. Biotrophic pathogens, on the other hand, establish a stable
relationship with living plant cells, obtaining nutrients by manipulating host cell
metabolism. Some pathogens, such as Ustilago maydis, the causal agent of corn
smut disease, are facultative biotrophs, being also capable of host-independent
growth. Others, such as the rust fungi, powdery mildews and downy mildews, are
obligate biotrophs, only growing and reproducing in association with a host
plant. In addition, a number of fungal and oomycete plant pathogens have an
intermediate life-style known as hemibiotrophy, where the pathogen initially
infects and propagates in living host tissue before switching to a necrotrophic
phase of disease development. Examples of hemibiotrophs include the rice blast
fungus, Magnaporthe oryzae, some Colletotrichum species and the oomycetes
Phytophthora infestans, P. sojae and P. nicotianae.
2 Infection Strategies
In order to infect plants, pathogens must overcome a range of highly effective
constitutive and induced, physical and chemical plant defences. In most cases, the
infection cycle involves four key stages – adhesion to the plant surface, penetration
Plant Infection by Biotrophic Fungal and Oomycete Pathogens
185
and colonisation of the plant, acquisition of nutrients and reproduction. Details of
the strategies followed in achieving these goals vary between different groups of
pathogens. A brief overview of the infection processes employed by the major
groups of biotrophic fungal and oomycete plant pathogens follows, with a particular
focus on penetration and colonisation.
2.1
Rust Fungi
Rust fungi belong to the Basidiomycetes and are some of the most successful
obligate biotrophs. A number have served as model systems for investigations of
plant–pathogen interactions. For example, genetic studies of Melampsora lini (flax
rust) formed the basis of the gene-for-gene hypothesis which states that every
resistance gene in a plant has a corresponding avirulence (Avr) gene in the pathogen
(Flor 1955). Molecular studies of Uromyces fabae (broad bean rust) have
contributed to our understanding of nutrient acquisition during biotrophic growth.
Due to their major contribution to disease development and propagation, dikaryotic
asexual rust uredospores have been the focus of most studies of rust infection
strategies (Fig. 1). When a uredospore lands on a plant, it adheres to the plant
surface initially through passive, hydrophobic interactions before secreting material
that forms an adhesive pad of glycoproteins and carbohydrates (Clement et al.
1993; Deising et al. 1996). Upon spore germination, the germ tube extends by tip
growth across the leaf surface. The detection of chemical signals on the plant
surface is no doubt a feature common to most, if not all fungi and oomycetes, but
hyphae of rust fungi also sense and respond to topographical signals from the
underlying surface. During this thigmotropic growth, hyphae of Puccinia species,
for example, extend in a direction that is perpendicular to the long axis of the leaf
epidermal cells (Read et al. 1992; Staples and Hoch 1997). When the hypha
encounters a guard cell, the hyphal apex swells and differentiates into an appressorium over the stomatal pore (Hoch and Staples 1991). Both thigmotropic growth
and appressorium formation can be induced on inert replicas of the leaf surface or
artificial gratings providing clear evidence that the inductive signals are associated
with topographical features of the underlying surface (Hoch et al. 1987). Entry of
the fungus into the plant leaf then occurs through the stomatal cavity via a
penetration hypha produced from the base of the appressorium.
On entering the leaf tissue, the penetration hypha expands to form a substomatal
vesicle in the substomatal cavity. From this vesicle, one or more primary infection
hyphae grow until they make contact with a host mesophyll cell whereupon they
differentiate to form haustorial mother cells (Heath and Skalamera 1997).
A penetration hypha is formed by the haustorial mother cell and grows through
the plant cell wall, invaginating the host plasma membrane as it expands and
differentiates into a haustorium (Harder and Chong 1991). Upon establishment of
haustoria, the fungus is no longer metabolically independent and enters a “parasitic/
186
P.H.P. Gan et al.
Fig. 1 Schematic representation of the rust fungal uredospore infection. Diagram design modified
from Voegele (2006). U uredospore, Ad adhesive, GT germ tube, Ap appressorium, PH penetration
hypha, SV substomatal vesicle, HMC haustorial mother cell, H haustorium, EHM extrahaustorial
membrane, GC stomatal guard cell, IH infection hyphae, C cuticle, UP uredospore pustule. Early
infection structures from the “penetration stage” are shown in dark brown, structures from the
“parasitic/biotrophic stage” are shown in yellow and structures from the “sporulation stage” are
shown in red
biotrophic phase” during which it obtains nutrients from its host (Deising et al.
1996; Voegele 2006).
2.2
Powdery Mildews
Powdery mildews are biotrophic ascomycetes that infect aerial parts of higher
plants. After landing on a plant, asexual conidia attach to the plant surface and
form one or, in the case of Blumeria graminis, two germ tubes (Fig. 2). In B.
graminis, growth of the first, the primary germ tube, is limited, while the second,
the appressorial germ tube, extends across the leaf, often growing along the
longitudinal grooves between adjacent epidermal cells (Carver et al. 1995). The
primary germ tube is believed to function in attaching the germling to the plant
surface, in obtaining water from the host, and in perception of chemical and
physical signals from the plant (Green et al. 2002). Signal transduction by the
primary germ tube influences subsequent germination and growth of the
appressorial germ tube. In response to inductive signals, the appressorial germ
tube swells at its tip and differentiates into a lobed appressorium which, via a
Plant Infection by Biotrophic Fungal and Oomycete Pathogens
187
Fig. 2 Schematic representation of Blumeria graminis infection. Ad adhesive, Ap appressorium,
PH penetration hypha, H haustorium, EHM extrahaustorial membrane, PGT primary germ tube,
Cm conidium, AGT appresorial germ tube, C cuticle. Early infection structures from the “penetration stage” are shown in dark brown and structures from the “parasitic/biotrophic stage” are shown
in yellow
penetration hypha, grows through the underlying host cuticle and cell wall (Green
et al. 2002). In contrast to rusts, powdery mildew appressoria form on the outer
periclinal wall of epidermal cells and directly penetrate the host cell wall. After
breaking through the cell wall, the penetration hypha develops into a haustorium
possessing finger-like projections that, as in rust infections, invaginate the plasma
membrane of the living host cell.
2.3
Magnaporthe oryzae
Magnaporthe oryzae is best known for the devastating blast disease it causes on rice
leaves (Talbot 2003) but it is also able to infect the roots of rice plants (Sesma and
Osbourn 2004; Marcel et al. 2010; Tucker et al. 2010). Depending on the host,
symptoms of M. oryzae infection can range from no visible symptoms to larger
necrotic lesions of different sizes indicating different proportions of necrotrophic
and biotrophic growth in fungal infection (Heath et al. 1991). Tissue-specific
differences in feeding strategies also exist. During foliar infections of rice,
M. oryzae is a hemibiotroph which infects and grows in living plant cells but,
after spreading to adjacent cells, it kills the initially infected cells (Wilson and
Talbot 2009; Kankanala et al. 2007). In rice roots, M. oryzae grows only
biotrophically (Wilson and Talbot 2009).
During foliar infection (Fig. 3), after landing on the plant, conidia of M. oryzae
release spore-tip mucilage which glues the spore to the plant surface (Hamer et al.
1988). The conidium germinates and the tip of the emergent germ tube swells and
differentiates into a dome-shaped appressorium. Cytoplasm flows from the
conidium into the appressorium as it enlarges and by the time the latter is mature,
the conidium is empty and collapsed (Braun and Howard 1994). The flat base of the
appressorium attaches tenaciously to the plant epidermis. The domed walls of the
cell become melanised, thereby greatly reducing permeability of the appressorial
wall and allowing passage of only water molecules (Howard et al. 1991).
188
P.H.P. Gan et al.
Fig. 3 Schematic representation of Magnaporthe oryzae foliar infection. Ad adhesive, GT germ
tube, Ap appressorium, PH penetration hypha, Cm conidium, IvH invasive hyphae, EIHM extrainvasive hyphal membrane, Pd plasmodesmata, BIC biotrophic interfacial complex, C cuticle.
Early infection structures from the “penetration stage” are shown in dark brown and structures
from the “parasitic/biotrophic stage” are shown in yellow
Concomitant accumulation of high concentrations of glycerol generates turgor
pressures as high as 8 MPa within the cell (Howard et al. 1991). A penetration
hypha containing actin microfilaments forms at the base of the appressorium and
breaches the underlying plant cell wall using both physical force and localised
enzymatic degradation of wall components (Howard et al. 1991). Within the
epidermal cell, M. oryzae forms bulbous invasive hyphae that are surrounded by
a host-derived extra-invasive hyphal membrane (Kankanala et al. 2007). Invasive
hyphae form a specialised structure termed the biotrophic interfacial complex
which is involved in secretion of some virulence factors (Khang et al. 2010).
Invasive hyphae grow into neighbouring cells via plasmodesmata in the intervening
cell wall and at this time, the previously infected cell dies (Kankanala et al. 2007).
During root infection, M. oryzae conidia germinate from both terminal cells
forming preinvasive hyphae and hyphopodia on the root surface (Tucker et al.
2010). Hyphopodia are hyphal swellings that, like appressoria, may aid host cell
penetration. Invasive hyphae are produced by both surface hyphae and hyphopodia
and breach the outer epidermal cell wall. The infection remains biotrophic with no
evidence of a switch to a necrotrophic phase (Marcel et al. 2010).
2.4
Colletotrichum Species
Most species of Colletotrichum are hemibiotrophs that grow biotrophically in living
plant cells for 1 to a few days before adopting a necrotrophic life-style. Like the
powdery mildews and M. oryzae, most Colletotrichum species penetrate the host
cuticle and cell wall of epidermal cells directly, rather than entering through a
stomatal aperture (Fig. 4; Willmer and Fricker 1996; Mendgen and Deising 1993).
Colletotrichum conidia germinate on the surface of a host and form a short germ
tube which responds to inductive signals on the plant surface and differentiates
into a dome-shaped appressorium. Structural and biochemical specialisation of
Plant Infection by Biotrophic Fungal and Oomycete Pathogens
189
Fig. 4 Schematic representation of Colletotrichum lindemuthum infection on a compatible host.
In other Colletotrichum species (e.g. Colletotrichum destructivum), only secondary, necrotrophic
hyphae spread from the initial invaded cell. Image design modified from Mendgen and Hahn
(2002). Ad adhesive, GT germ tube, Ap appressorium, PH penetration hypha, Cm conidium,
V vesicle, PrH primary hyphae, SdH secondary hyphae, C cuticle. Early infection structures
from the “penetration stage” are shown in dark brown and structures from the “parasitic/biotrophic
stage” are shown in yellow
Colletotrichum appressoria are similar to those of M. oryzae – the appressorial cell
wall becomes impregnated with melanin and the cell accumulates high
concentrations of osmotically active compounds (Mendgen and Deising 1993). In
C. graminicola, the resultant appressorial turgor pressure has been calculated to
exceed 5.2 MPa (Bechinger et al. 1999). As in M. oryzae, the appressorial penetration hypha breaches the underlying epidermal cell wall, probably by using a
combination of mechanical force and enzymatic cell wall degradation (M€unch
et al. 2008). Within the epidermal cells, the tip of the penetration hypha swells to
form a vesicle and broad primary intracellular hyphae that remain separated from
the host cytoplasm by an interstitial matrix and an invaginated host-derived plasma
membrane. This relationship lasts for 1–2 days after which the plant plasma
membrane is degraded and the infected cell dies. In Colletotrichum
lindemuthianum, this transient biotrophy is maintained in newly invaded cells but
2–3 days after the onset of infection, narrow secondary hyphae form and breach the
plant plasma membrane, killing the host cells. The pathogen thus enters its
necrotrophic phase during which secretion of a diversity of cell wall degrading
enzymes leads to widespread cell wall digestion (M€unch et al. 2008). In
Colletotrichum destructivum, only secondary necrotrophic hyphae spread from
the initial invaded cell (Perfect and Green 2001). In Colletotrichum capsici, infection hyphae initially grow between the cuticle and cell wall of the epidermal cells
before killing and colonising the plant tissues (Mendgen and Hahn 2002).
2.5
Oomycetes
Oomycete plant pathogens include obligate biotrophs (e.g. the downy mildews),
hemibiotrophs (e.g. P. infestans) and necrotrophs (e.g. Phytophthora cinnamomi).
Some are foliar pathogens; others infect plant roots. Hyphae of oomycete species
190
P.H.P. Gan et al.
may penetrate the plant surface via stomata, along the anticlinal wall between
adjacent epidermal cells or directly through the outer periclinal cell wall of the
epidermis. Biotrophic and hemibiotrophic species form simple, spherical or pyriform haustoria (Enkerli et al. 1997). In hemibiotrophic Phytophthora species, tissue
colonisation is typically through intercellular hyphal growth with haustoria
differentiating within plant cells contacted by the intercellular hyphae (Enkerli
et al. 1997).
3 Structure and Function of Fungal and Oomycete Haustoria
Haustoria formed by biotrophic or hemibiotrophic rusts, powdery mildews and
oomycetes are central to the development of a stable biotrophic relationship
between pathogen and plant and are key sites for the absorption of the plant
nutrients that are essential for pathogen growth, development and reproduction.
For their function, haustoria depend on highly regulated molecular exchange across
the haustorium–plant boundary. Before nutrients begin to move from plant to
pathogen across this interface, the pathogen must send a likely plethora of signals
into the infected plant cell, molecules that not only orchestrate host cell structure
and metabolism but also suppress the plant cell’s defence response. Given the
importance of communication across this haustorium–plant interface, it is not
surprising that the area of this boundary is often increased by the development, in
the rusts, of up to five lobes (Littlefield 1972) or, in the powdery mildews, of
multiple finger-like projections (Gil and Gay 1977) extending from the main body
of the haustorium. There is still much to learn about the structure and function of the
haustorium–plant interface but integrated cellular and molecular studies are beginning to reveal details of its molecular specialisations.
The composition of both the haustorial cell wall and plasma membrane may
differ from that of the cell wall and plasma membrane of hyphae. In flax rust, for
example, monoclonal antibodies identify a component of the haustorial wall not
found in hyphal walls of this species (Murdoch and Hardham 1998). The same
monoclonal antibodies do not react with haustoria from maize rust (Puccinia
sorghi) or wheat leaf rust (Puccinia recondita), indicating that the haustorialspecific component may also be species-specific (Murdoch and Hardham 1998).
Differentiation of haustorial membranes has been demonstrated in the bean rust
fungus (see below) and in the hemibiotrophic oomycete, P. infestans (Avrova et al.
2008). In P. infestans, haustorial membranes, but not membranes of intercellular
hyphae, contain the membrane protein, Pihmp1, a protein required for haustoria
formation (Avrova et al. 2008).
Haustoria are surrounded by an amorphous, gel-like compartment known as the
extrahaustorial matrix, across which molecular exchange between pathogen and
plant must occur as shown in Fig. 5 (O’Connell 1987; Hahn and Mendgen 1992;
Mendgen and Hahn 2002). The matrix consists of a mixture of components,
primarily carbohydrates and glycoproteins, derived from both the fungus and the
Plant Infection by Biotrophic Fungal and Oomycete Pathogens
191
Fig. 5 The plant-haustorium interface. In haustoria-forming biotrophic fungi, the haustorial
mother cell (HMC) forms a haustorium (H) surrounded by the extrahaustorial matrix (EHMX),
which is a discrete compartment sealed by a neckband (N). Upper inset: A proton gradient
generated by ATPases (purple oval) on the haustorial membrane (HM) may facilitate uptake of
metabolites into the fungus through proton-driven symporters (green oval). Lower inset: Effectors
(blue dots) may cross the extrahaustorial membrane (EHM), a modified host plasma membrane
that invaginates around the haustorium, into the host cytoplasm, where they may interact with
target proteins, thus leading to host manipulation or resulting in recognition by cognate resistance
proteins and the mounting of a defence response. Effectors may also be secreted into the apoplast.
PM plant plasma membrane, CW plant cell wall, HCW haustorial cell wall
plant (Harder and Chong 1991). The extrahaustorial matrix is delimited on
the pathogen side by the haustorial cell wall and on the host side by an extension
of the host plasma membrane known as the extrahaustorial membrane (Harder and
Chong 1991). In rust and powdery mildew infections, the extrahaustorial matrix is
sealed off from the apoplast by a structure known as the neckband which is thought
to act as a selectively permeable barrier, like the Casparian strip at the endodermis
of plant roots (Heath 1976; Harder and Chong 1991). Haustoria of hemibiotrophic
fungi and oomycetes appear to be less differentiated than those of the rusts and
powdery mildews and generally lack neckbands, although some exceptions exist, as
in the case of the oomycete Albugo candida which possesses a simple neckband
(Soylu 2004; Perfect and Green 2001). In infections by some rust fungi, such as
Puccinia hemerocallidis and Puccinia striiformis, tubular beaded elements contiguous with the extrahaustorial matrix extend from the extrahaustorial membrane into
the host cytoplasm, perhaps again serving to increase the surface area of this
interface (Mendgen et al. 1991; Mims et al. 2002, 2003). These structures were
not observed at the host–haustorial interface during P. recondita or Uromyces
appendiculatus rust infection (Mendgen et al. 1991). Tubular extensions into the
cytoplasm from the extrahaustorial membrane also occur during infections by the
oomycete A. candida (Baka 2008).
Although contiguous with the plant plasma membrane, from which it is thought
to be derived, the extrahaustorial membrane exhibits distinct molecular properties
192
P.H.P. Gan et al.
(Hardham 2007). Differences include a reduced glycolipid content, differential
staining by phosphotungstic acid and a reduced level of ATPase activity compared to the rest of the host plasma membrane (Baka et al. 1995; Perfect and
Green 2001). The extrahaustorial membrane formed during infection of pea
plants by the pea powdery mildew, Erysiphe pisi, contains a large (200 kDa)
glycoprotein that does not occur elsewhere in the plasma membrane of infected
cells (Micali et al. 2008; Roberts et al. 1993). Similarly, the plant resistance
protein RPW8.2 is specifically targeted to the extrahaustorial membrane of
Arabidopsis leaf cells infected by powdery mildew, Golovinomyces orontii
(Micali et al. 2010). During infections of Arabidopsis by G. orontii and another
powdery mildew, Erysiphe cichoracearum, host plasma membrane proteins
are specifically excluded from the extrahaustorial membrane (Koh et al. 2005;
Micali et al. 2010).
Haustoria have been difficult to analyse at a molecular level because they are only
formed in planta. However, methods to isolate haustoria either by differential
centrifugation or affinity purification have been developed (Gil and Gay 1977;
Tiburzy et al. 1992; Hahn and Mendgen 1992; Micali et al. 2010) and in recent
years have facilitated studies of haustorial proteins and transcriptomes (Hahn et al.
1997; Murdoch and Hardham 1998; Catanzariti et al. 2006; Micali et al. 2010). Early
studies led to the proposal that haustoria were specialised feeding structures that
played a key role in nutrient acquisition, but definitive evidence was difficult to
obtain. Strong evidence to support this hypothesis has come from transcriptome
analyses of genes that are preferentially expressed in haustoria of the bean rust
fungus, U. fabae (Hahn and Mendgen 1997). This study identified two cDNA
sequences, AAT1 and AAT2, which have homology to genes encoding amino acid
transporters (Hahn and Mendgen 1997). AAT1 is a broad-specificity amino acid
transporter (Struck et al. 2002). AAT2 was shown by immunolocalisation to occur
specifically in the haustorial membrane, although no amino acid transport activity
has yet been described for this protein (Mendgen et al. 2000). Further evidence for a
role of U. fabae haustoria in nutrient acquisition comes from studies of a haustorially
expressed sugar uptake transporter, HXT1p, which transports monosaccharides
through a proton symport process when expressed in Xenopus oocytes (Voegele
et al. 2001; Voegele and Mendgen 2003). Given that ATPases, which are required
for the pumping of protons across membranes, are reduced on the extrahaustorial
membrane but enriched on the fungal haustorial membrane (Baka et al. 1995; Struck
et al. 1996, 1998), this finding suggests that during infection, a proton gradient is
generated between the extrahaustorial matrix and the haustorium that is used to drive
active transport of nutrients towards the pathogen (Fig. 5 inset; Szabo and Bushnell
2001; Baka et al. 1995; Aist and Bushnell 1991).
In addition to this role in nutrient acquisition, it has also emerged that the
secretion of pathogen proteins, known as effectors (described in greater detail
below), from haustoria is important in establishing biotrophy by allowing manipulation of host metabolism and defence responses. In agreement with this scenario,
compatibility between the host and the pathogen appears to be determined at the
onset of haustorial development since spore germination, germ tube growth,
Plant Infection by Biotrophic Fungal and Oomycete Pathogens
193
appressorium formation and growth of primary infection hyphae develop on a
similar time scale in both susceptible and resistant host plants (Dickinson and
Lucas 1982).
4 Intracellular Infection Hyphae
Some biotrophic or hemibiotrophic fungal pathogens, such as M. oryzae, U. maydis
and Colletotrichum species, do not produce haustoria but instead form specialised
intracellular hyphae that may serve the same or a similar function (Mendgen and Hahn
2002; O’Connell and Panstruga 2006). Like haustoria, these intracellular infection
hyphae remain separated from the host cytoplasm during biotrophic growth. Intracellular hyphae of M. oryzae, for example, are surrounded by a host-derived plasma
membrane, termed the extra-invasive hyphal membrane, and an extra-invasive hyphal
matrix (Kankanala et al. 2007). Although no neckband structure has been detected
which might seal the space between the fungus and the host membrane, the extrainvasive hyphal matrix appears to be distinct from the plant apoplast since addition of
the lipid-binding dye FM4-64 to the apoplast leads to the incorporation of the dye into
the plasma membrane of plant cells but not of the intracellular hyphae (Kankanala
et al. 2007). In hemibiotrophic pathogens, the switch to necrotrophy is associated
with distintegration of the host plasma membrane around the intracellular fungal
structures, suggesting that the zone of separation between host and pathogen
is important in maintaining a biotrophic relationship (Voegele et al. 2009).
5 Host–Pathogen Specificity
As described in the previous sections, biotrophic and hemibiotrophic fungi and
oomycetes are able to grow within living host cells and tissues. In order to maintain
this intimate association, the pathogen alters and manipulates the host metabolism
extensively. Among the changes that have been documented in host plants during
infection are movement of the host nucleus and endomembrane system to surround
the extrahaustorial membrane; a redistribution of carbohydrates into infected
leaves; and an increase in chlorophyll and photosynthetic activity, leading to the
“green island effect” (Scholes and Farrar 1985; Coffey 1972; Leckie et al. 1995;
Voegele et al. 2009). These events occur during a compatible interaction but in an
incompatible interaction plant cells are able to sense pathogen invasion and react
rapidly by mounting a multi-faceted defence response.
Plants are able to detect both chemical and physical cues associated with
potential pathogens. Currently, plants are seen to have two major categories of
defence response. The first, often termed basal resistance, occurs at the onset of
infection and is triggered by recognition of conserved pathogen-derived molecules,
such as chitin fragments from fungal cell walls or proteins or glucans from
194
P.H.P. Gan et al.
oomycete cell walls (Cosio et al. 1996; Brunner et al. 2002; Kishimoto et al. 2010).
These molecules are collectively referred to as elicitors or pathogen- or microbeassociated molecular patterns (PAMPs or MAMPs) and are recognised by plant
pattern recognition receptors (PRRs) (Boller and Felix 2009). A number of PRRs
have been identified. They include the rice CEBiP receptor that recognises fungal
chitin (Kaku et al. 2006) and a 75 kDa protein that binds Phytophthora cell wall bglucans (Mithofer et al. 2000). Detection of PAMPs by PRRs results in a localised
basal defence response known as PAMP-triggered immunity (PTI) (Jones and
Dangl 2006).
One of the earliest responses observed during PTI is the aggregation of cytoplasm in the host cell adjacent to the pathogen appressorium or invading hypha
(Freytag et al. 1994). This is accompanied by reorganisation of the actin cytoskeleton and endomembrane system to focus on the penetration site (Takemoto et al.
2003), and reinforcement of the cell wall by cell wall appositions (papillae) (Aist
1976). Wall apposition formation involves localised deposition of structural wall
components and anti-microbial compounds and is achieved by the actin-based
targeting of secretion. During infection of barley by the powdery mildew
B. graminis, for example, multivesicular bodies and vesicles containing hydrogen
peroxide and phenolics, accumulate in the host cell at the site of penetration
(An et al. 2006). PTI-associated defence responses also include the generation of
reactive oxygen species (ROS) and the induction of defence-related genes which
may inhibit further growth of the invader (Torres 2010; Schwessinger and Zipfel
2008). As demonstrated during the interaction of Uromyces vignae with the nonhost broad bean plant, clathrin-mediated endocytosis in the plant is active at this
early stage of infection and may function to recycle activated defence-related
receptors from the plant surface (Xu and Mendgen 1994; Leborgne-Castel et al.
2010). Similar reorganisation of the actin and endomembrane systems can be
triggered by mechanical stimuli but the extent of the defence response evoked
has not yet been determined (Gus-Mayer et al. 1998; Hardham et al. 2008).
Virulent (adapted) pathogens are able to avoid or suppress PTI. Avoidance
mechanisms include the masking or removal of potential PAMPs. Pre-infection
by an adapted pathogen has been shown to permit subsequent growth by non-host
pathogens, indicating that a general suppression of basal resistance has occurred
(Fernandez and Heath 1991). Adapted pathogens can suppress PTI through the
synthesis and secretion of specific effector proteins (Jones and Dangl 2006), as
discussed in detail below.
Plants have, in turn, evolved resistance proteins that have the ability to recognise
pathogen effectors which, in an incompatible interaction, are often called Avr
proteins. Plant resistance proteins have a nucleotide binding (NB) domain and a
C-terminal leucine-rich repeat (LRR) region (Dodds and Rathjen 2010). In addition, many resistance proteins also have an N-terminal Toll, interleukin-1 receptor
domain or a coiled-coil domain (Dodds and Rathjen 2010). These proteins have
structural similarity to proteins that have been found to be important for immunity
in animal systems, namely, the Toll receptors in Drosophila and Toll-like receptors
in mammals (Staskawicz et al. 2001). Collectively NB-LRR proteins confer
Plant Infection by Biotrophic Fungal and Oomycete Pathogens
195
resistance against various pathogens including fungi, oomycetes, bacteria, viruses
and insects (Dodds and Rathjen 2010). Recognition of an Avr protein by a corresponding resistance protein results in effector-triggered immunity (ETI), which
typically culminates in a form of programmed cell death known as the hypersensitive response (HR) (Dodds and Rathjen 2010; Jones and Dangl 2006). In a limited
number of cases, both the plant resistance protein and the corresponding pathogen
Avr protein have been identified. For example, genes encoding the M and L6
resistance proteins of flax plants and their corresponding Avr proteins, AvrM and
AvrL567, of flax rust, have been cloned and the encoded resistance proteins and
Avr proteins shown to interact directly (Catanzariti et al. 2010; Lawrence et al.
1995; Anderson et al. 1997; Dodds et al. 2004, 2006; Catanzariti et al. 2006).
6 Effectors
The discovery of pathogen effectors that can “manipulate host cell structure and
function, thereby facilitating infection (virulence factors or toxins) and/or triggering defence responses (avirulence factors or elicitors)” (Kamoun 2006) has led to
intense interest and effector-focused research. Although initially identified slowly
and laboriously by positional cloning (Orbach et al. 2000; Dodds et al. 2004; Ridout
et al. 2006; Gout et al. 2006), advances in DNA sequencing and bioinformatic
technologies now facilitate the prediction of suites of candidate effectors in entire
phytopathogen genomes (Torto et al. 2003; Choi et al. 2010). Typically, criteria
used to interrogate pathogen genomes include identifying sequences encoding
small proteins which are under diversifying selection and which have an N-terminal
signal peptide (Mueller et al. 2008; Dean et al. 2005; Kamper et al. 2006; Haas et al.
2009). These search strategies have also been applied to cDNA libraries and EST
(Expressed Sequence Tags) databases enriched with sequences from the biotrophic
phase of pathogenic biotrophs or from the host–pathogen interface. This approach
has, for example, resulted in the identification of effectors AvrM, AvrP4 and
AvrP123 from M. lini (Catanzariti et al. 2006).
Many important insights into effector biology have been acquired by studying
the sequences of predicted effectors. Analysis of the sequences of known
Phytophthora effectors has, for example, led to identification of a conserved
N-terminal RXLR dEER motif in these proteins (RXLR-X5-21-dEER; where
upper case letters represent consensus amino acids in 10 out of 11 sequences and
lowercase letters represent consensus amino acids in more than half of the genes)
(Rehmany et al. 2005; Birch et al. 2006). Using this motif as a search criterion,
150–600 candidate effectors have been identified in the genomes of Phytophthora
sojae and Phytophthora ramorum, the exact number obtained depending on the
stringencies of the search settings, such as whether or not the dEER motif is also
included and if hidden Markov Model statistics are applied (Win et al. 2007; Tyler
et al. 2006). Other motifs that have been identified by sequence comparisons of
potential Phytophthora effectors include the W-, L- and Y-motifs in the C-terminal
196
P.H.P. Gan et al.
portions of some RXLR effectors (Jiang et al. 2008). The W-motif is required for
the HR mediated by the interaction between the Rpi-blb1 resistance protein and
ipiO effectors (Champouret et al. 2009) and both W- and Y-motifs are required for
suppression of programmed cell death by Avr1b (Dou et al. 2008a).
Another family of Phytophthora effectors known as Crinklers was identified by
analysis of ESTs encoding secreted sequences (Torto et al. 2003). Members of this
gene family in P. infestans, P. sojae and P. ramorum cause necrosis in host plants
and contain a conserved LXLFLAK motif at the N-terminus (Haas et al. 2009;
Win et al. 2007). Similar analyses of EST sequences isolated from barley epidermal
cells containing B. graminis powdery mildew haustoria identified a Y/F/WxC-motif
in small secreted proteins encoded by this fungus (Godfrey et al. 2010). However,
not all pathogen effectors have such readily recognisable motifs and, to date, no
highly conserved motif has been associated with other fungal effector sequences
[e.g. in flax rust (Rafiqi et al. 2010) and in poplar rust (Joly et al. 2010)]. In the
majority of cases, the function and biological significance of the putative effectors
identified by bioinformatics remain to be determined. To this end, several highthroughput techniques for functional screening of potential effectors have been
developed, including screening for phenotypes arising from expression of proteins
in host plants transformed by Agrobacterium (Torto et al. 2003; Oh et al. 2009).
7 Importance of Effectors During Infection
The large number of effectors predicted from sequence analyses hints at the complex
nature of pathogen–host interactions. While it is possible that not all of the predicted
effectors actually function as such, cohorts of effectors may be required at different
stages of infection. For example, the deletion of a potential virulence cluster from the
genome of U. maydis leads to defects in tumour and spore formation, but does not
affect penetration or hyphal growth of the fungus within plant tissue (Kamper et al.
2006; D€
oehlemann 2011). Different subsets of effectors may also be required
according to the age and type of host tissue being infected (Skibbe et al. 2010).
Multiple variants of a given effector exist [(e.g. flax rust has six AvrM sequence
variants (Catanzariti et al. 2006)], due perhaps to evolutionary selective pressure to
evade recognition by host resistance proteins and to retain virulence. Multiple
sequence variants may also inhibit effector recognition. In the case of the multigene
P. infestans ipiO family, normal recognition of the IPI-O1 variant by the RBresistance protein is impaired in the presence of the IPI-O4 variant, which is not
recognised by the RB-resistance protein (Halterman et al. 2010).
Redundant or non-essential effector function may limit the value of knockout or
silencing studies, as illustrated by studies in which the flax rust AvrL567 gene was
silenced with no apparent effect on pathogen virulence (Lawrence et al. 2010).
However, a number of effectors have been shown to interfere with plant basal
defence responses. In C. fulvum, two effectors inhibit chitin recognition and
ensuing induction of PTI. The Avr4 effector protein binds to chitin in the fungal
cell wall possibly protecting it from host chitinases (van den Burg et al. 2006), thus
Plant Infection by Biotrophic Fungal and Oomycete Pathogens
197
maintaining fungal cell wall integrity and inhibiting release of the chitin-derived
PAMPs that trigger PTI. A second C. fulvum effector, Ecp6, binds to chitin
fragments and effectively competes with host PAMP recognition receptors for
chitin binding, again preventing PTI induction (de Jonge et al. 2010). Some
effectors appear to inhibit downstream PTI signalling processes. The P. infestans
Avr3a effector, for example, suppresses cell death induced by the elicitor INF1 in
plants that lack the R3a resistance protein (Bos et al. 2006, 2009). Further, during
Fusarium oxysporum infection of tomato, the effector Avr1 inhibits HR induced by
Avr2 and Avr3 (Houterman et al. 2008).
A number of effectors are predicted to target host enzymes that may be involved
in plant defence. These include the EPI1 Kazal-like protease inhibitor from
P. infestans which inhibits the P69B protease, the EPIC2B effector which inhibits
host papain proteases PIP1 and RCR3, the C. fulvum Avr2 effector which also
inhibits RCR3, and the P. infestans glucanase inhibitors GIP1 and GIP2 (Tian et al.
2005, 2007; Song et al. 2009; Rooney et al. 2005; van Esse et al. 2008; Shabab et al.
2008; Rose et al. 2002; Damasceno et al. 2008; Hein et al. 2009). The flax rust
effector, AvrP123, has also been shown to have homology to Kazal-like protease
inhibitors although the biological significance of this has not yet been demonstrated
(Catanzariti et al. 2006). According to Kamoun’s (2006) definition, the great
diversity of cell wall degrading enzymes produced by pathogens constitute extracellular effectors (Hardham and Cahill 2010).
In many cases, the functions of putative effectors are yet to be determined,
although many identified effectors are Avr proteins and are thus known to trigger
the resistance response when detected by a corresponding resistance protein.
Interestingly, mutational analysis of Avr3a indicates that the Avr and virulence
functions of the effector can be separated, since mutants that are recognised by the
R3a resistance protein but which do not suppress INF1-induced cell death have
been isolated (Bos et al. 2009). In the flax-flax rust system, yeast two-hybrid assays
show that avirulent but not virulent forms of the AvrL567 and AvrM effectors
interact directly with the L6 and M resistance proteins, respectively (Dodds et al.
2006; Catanzariti et al. 2010). AvrPita from M. oryzae was also shown to bind the
cytoplasmic Pi-ta resistance protein from rice (Jia et al. 2000).
8 Delivery of Effectors into Host Cells
While many of the fungal and oomycete effectors discussed in the previous section
are likely to function in the plant apoplast, the direct interaction between Avr
proteins and their cognate resistance proteins known to occur within the plant
cytoplasm suggests that pathogen effectors must also function in the host
cytoplasm.
Early evidence for delivery of fungal proteins into host plant cells came from
immunolocalisation of UfRTP1, a haustorially expressed glycoprotein of unknown
function secreted by the broad bean rust (Kemen et al. 2005). UfRTP1 was shown to
198
P.H.P. Gan et al.
Fig. 6 Localisation of the flax rust effector AvrM in planta. (a–c) Immunofluorescence
localisation of AvrM in leaves infected with the flax rust fungus. (a) DIC image of fluorescence
micrographs showing labelling of haustoria (b) and AvrM (c). The AvrM signal is strong at and
around haustoria (arrowheads) but is also detected in the cytoplasm of invaded plant cells. (d)
Immunogold labelling of AvrM in flax rust-infected leaves. The majority of AvrM labelling is
detected in the extrahaustorial matrix (EHMX). However, lower intensity signal was also detected
within the plant cytoplasm (Cy). HCW haustorial cell wall, EHM extrahaustorial membrane
occur in the extrahaustorial matrix and within the host cell during infection (Kemen
et al. 2005). UfRTP1 contains a nuclear localisation signal and accumulates in the
host nucleus during infection (Kemen et al. 2005). More recently, immunolocalisation experiments with antibodies raised against AvrM have provided direct
evidence that this effector is secreted by fungal hyphae and haustoria during
infection and enters the cytoplasm of host cells containing one or more haustoria
(Fig. 6; Rafiqi et al. 2010). Localisation of AvrM within the plant cytoplasm is
consistent with the recent demonstration of the M resistance protein within the host
cytosol (D. Takemoto and D. A. Jones, unpublished observations). Agrobacteriummediated transient expression of flax rust effectors AvrL567 and AvrM lacking their
signal peptides also triggers HR in plants that contain the corresponding resistance
proteins, providing further evidence of the function of these proteins within the plant
cytoplasm (Dodds et al. 2004; Catanzariti et al. 2006). Expression of wild-type
(secreted) AvrL567 and AvrM fused to fluorescent proteins via Agrobacteriummediated transformation demonstrates that the predicted signal peptides of the
fungal effectors are functional in directing protein secretion and that after their
secretion from the plant cell, the effectors can re-enter the cytoplasm of both tobacco
and flax plant cells (Rafiqi et al. 2010). These experiments provide evidence that
uptake of fungal effectors into host plant cells is pathogen-independent, requiring
host factors but not pathogen components (Rafiqi et al. 2010). The conservation of
Plant Infection by Biotrophic Fungal and Oomycete Pathogens
199
the uptake mechanism between different plant species is consistent with the findings
of a study in which purified AvrL567-GFP protein was internalised into soybean
root cells and into human A549 cells (Kale et al. 2010).
Localisation of truncation and deletion mutants of AvrM and AvrL567 effector
proteins have identified N-terminal regions of less than 31 amino acids that are
sufficient to mediate re-uptake of fluorescently tagged protein into host cells (Rafiqi
et al. 2010). The primary amino acid sequences of these putative uptake domains
are not conserved between the two effectors (Rafiqi et al. 2010). Transient expression of cytoplasmic effectors AvrP123 and AvrP4 from M. lini and Avra10 and
Avrk1 from B. graminis in plants with the corresponding R genes also elicits HR,
indicating that delivery of effectors to the plant cytoplasm where they are
recognised as part of ETI is a common phenomenon in biotrophic fungal pathogens
(Catanzariti et al. 2006; Ridout et al. 2006; Barrett et al. 2009).
Many oomycete effectors also enter the host cytoplasm during infection, but in
contrast to the situation with fungal pathogens, studies of the oomycetes have
identified two domains involved in transport into the plant cell cytoplasm. In
haustoria-forming oomycetes, the N-terminal RXLR motif mediates effector
uptake into plant cells (Whisson et al. 2007; Dou et al. 2008b). In oomycete
effectors in the Crinkler family, the conserved N-terminal LXLFLAK motif downstream of the signal peptide is required for targeting of Crinkler proteins into host
cells (Schornack et al. 2010; Grenville-Briggs et al. 2010). Crinkler effectors are
conserved more widely among oomycetes than the RXLR effectors and some
necrotrophic oomycetes, such as the legume rot pathogen Aphanomyces euteiches,
that do not form haustoria possess Crinkler but not RXLR effectors (Schornack
et al. 2010). A recent proteomics study has also identified a cell wall-associated
Crinkler protein, suggesting that these effectors may function in either the plant
apoplast or cytoplasm (Grenville-Briggs et al. 2010). This effector contains the
LXLFLAK motif and it remains possible that it might function both intracellularly
as well as extracellularly.
As with the flax rust effectors, the mechanism of RXLR effector uptake into
host cells is pathogen-independent (Dou et al. 2008b). Purified Avr1b-GFP from
P. sojae is taken up by soybean root and human A549 cells (Dou et al. 2008b; Kale
et al. 2010) and purified SpHTP1-GFP from Saprolegnia parasitica is taken up by
host fish cells (van West et al. 2010). These results indicate that the mechanism of
RXLR entry is not only pathogen-independent but exploits an uptake pathway that
is widely conserved among eukaryotic cells.
In M. oryzae, which lacks haustoria but forms intracellular invasive hyphae,
early studies showed that AvrPita, the effector that interacts directly with the rice
cytoplasmic Pi-ta resistance protein, triggered a HR when expressed in the cytoplasm of plants expressing Pi-ta via particle bombardment, suggesting that AvrPita
enters host cells where it is recognised by the resistance protein (Jia et al. 2000).
Recent studies tracking the delivery of M. oryzae effectors into host cells have
revealed that a number of pathogen effectors, including AvrPita, PWL1, PWL2 and
BAS1 are secreted from the biotrophic interfacial complex into the plant cell
cytoplasm (Figs. 1–3; Khang et al. 2010). Initial secretion into the biotrophic
200
P.H.P. Gan et al.
interfacial complex is directed by the signal peptide and/or part of the promoter
encoding sequence since these regions of AvrPita, PWL1 and PWL2 are sufficient
to direct fluorescently tagged proteins into the biotrophic interfacial complex
(Khang et al. 2010). The role of the biotrophic interfacial complex and the mechanism underlying effector uptake across the host membrane has yet to be elucidated.
Passage of fungal and oomycete effectors into the plant cell cytoplasm is a
crucial aspect of pathogenicity and elucidation of the mechanism of effector uptake
is clearly an important step in understanding the infection process. In contrast to the
Type III secretion system of Gram-negative bacteria (Cornelis 2006; Alfano and
Collmer 2004; Galan and Wolf-Watz 2006) or the PEXEL protein export system of
the Plasmodium malarial parasites (Boddey et al. 2010; de Koning-Ward et al.
2009), uptake of fungal and oomycete effectors does not require participation of
any other pathogen-encoded molecules. Uptake of fungal and oomycete effectors
depends solely on plant-encoded components.
Several mechanisms of pathogen-independent protein uptake into the cytosol of
host cells have been proposed, as summarised in Fig. 7. These can be broadly
categorised into (1) endocytic uptake, (2) transport through a transmembrane
transporter and (3) direct membrane penetration (Drin et al. 2003; Goldberg and
Cowman 2010; Voegele et al. 2009). After endocytosis, material may be released
via retrotranslocation through the Golgi apparatus and endoplasmic reticulum (ER)
into the cytoplasm (route 1a) or directly from the endosomes (route 1b). Retrotranslocation is normally associated with cellular protein quality control, a process in
which misfolded proteins are transported from the ER into the cytosol where they
are targeted for proteosomal degradation. However, some pathogen proteins, such
as the Shiga and ricin toxins, are able to subvert this process for delivery into the
host cytosol (Magzoub et al. 2005; Sandvig and van Deurs 1996; Sandvig and van
Deurs 2002).
Multiple forms of endocytosis are known to occur in animal cells, with the main
two mechanisms involving clathrin-mediated and lipid raft-associated pathways
(Doherty and McMahon 2009; Mayor and Pagano 2007). Plant genomes include
genes encoding a number of proteins that function in clathrin-mediated endocytosis
(Barth and Holstein 2004; Holstein and Oliviusson 2005) and clathrin-mediated
uptake has been shown to participate in endocytosis of the PIN auxin efflux carrier
(Dhonukshe et al. 2007). During infection of broad bean by U. fabae, coated
vesicles are seen in the plant cytoplasm surrounding the haustorium, suggesting
that clathrin-dependent endocytosis is taking place (Xu and Mendgen 1994),
however, whether or not endocytosis might be internalising U. fabae effectors is
not known. Recent inhibitor studies did not give any evidence to suggest that uptake
of the flax rust effector AvrL567 and the P. infestans effector Avr1b into human
A549 cells requires clathrin-dependent receptor-mediated endocytosis (Kale et al.
2010). However, immunogold localisation of the flax rust effector AvrM in high
pressure-frozen and freeze-substituted material does provide evidence of vesiclemediated uptake of AvrM (P.H.P. Gan and A.R. Hardham, unpublished
observations). In animal cells, some forms of lipid raft-associated endocytosis
involve caveolin or flotillin proteins. Although there is no evidence of caveolin
Plant Infection by Biotrophic Fungal and Oomycete Pathogens
201
Fig. 7 Possible routes for effector entry. Effectors (red dots) may enter plant cells in the absence
of the pathogen via endocytic uptake (1a + b), where escape from endosomes into the cytosol may
occur via retrotranslocation through the Golgi apparatus and the endoplasmic reticulum (ER) (1a)
or via endosomal escape (1b); transfer through a transmembrane transporter (2) and/or through
direct membrane penetration (3). Figure design modified from Voegele et al. (2009). EHMX
extrahaustorial matrix
in plant genomes (Aniento and Robinson 2005), flotillins have been localised to
lipid rafts in the plasma membrane in Arabidopsis and M. truncatula (Haney and
Long 2010; Borner et al. 2005). In M. truncatula, establishment of nodules by the
bacteria Sinorhizobium meliloti during symbiosis requires two flotillins, possibly
for the trafficking of signals co-ordinating early symbiotic events or regulating
nodulation (Haney and Long 2010).
The possible role of lipid rafts in fungal and oomycete effector uptake has
recently been explored as part of a study of effector binding to membrane lipids
(Kale et al. 2010). The proposal arising from this investigation is that fungal and
oomycete effector entry into host cells is mediated by binding to the lipid
phosphatidylinositol-3-phosphate (PtdIns(3)P) on the extracellular leaflet of the
lipid bilayer of the host plasma membrane (Kale et al. 2010). This study presents
evidence that oomycete effectors bind to PtdIns(3)P via the RXLR motif and that
this interaction is required for effector uptake (Kale et al. 2010). Phosphotidylinositides constitute a small but important group of phospholipids within cells.
They are involved in signalling processes and contribute to spatial regulation of
cellular organisation through enrichment of different subsets of phosphotidylinositides in specific organelle membranes or membrane domains (Heilmann
2009). Kale et al. (2010) suggest that other fungal effectors, including AvrL567
from M. lini and AvrLm6 and AvrLm2 from L. maculans, may also enter host
cells by PtdIns(3)P-binding. However, although AvrM was observed to bind to
PtdIns(3)P, studies of GST fusions of deletion AvrM mutants have shown that
polypeptides that are taken up in planta do not necessarily exhibit PtdIns(3)P
binding activity (Gan et al. 2010), indicating that PtdIns(3)P-binding is not required
for AvrM entry into host cells.
202
P.H.P. Gan et al.
Another fungal protein that internalises into plant cells in the absence of the
pathogen is the PtrToxA toxin from the causal agent of wheat tan spot,
Pyrenophora tritici-repentis (Manning and Ciuffetti 2005). Toxin entry is dependent on the presence of an RGD vitronectin-like motif (Manning et al. 2008). In
animal cells, the RGD trimer mediates binding of extracellular matrix proteins to
integrins in the plasma membrane, and integrins are known to be targets of
mammalian pathogens which internalise into host cells via a form of clathrinmediated uptake (Isberg and Tran Van Nhieu 1994). Uptake of PtrToxA occurs in
ToxA-sensitive but not ToxA-insensitive host mesophyll cells and the protein
accumulates close to the infiltration site, suggesting interaction with a high-affinity
receptor on the surface of the host cell and internalisation via a form of receptormediated endocytosis (Manning and Ciuffetti 2005). The RGD motif is also present
in the P. infestans IPI-O effector (which also contains an RXLR motif) and IPI-O
has been shown to compete with other RGD-containing peptides in binding to the
plant plasma membrane (Senchou et al. 2004). However, this motif is absent from
the AvrM and AvrL567 minimum uptake regions identified by Rafiqi et al. (2010),
indicating that binding to integrins may not be the mechanism of host cell entry for
these fungal effectors.
Another category of proteins that are able to enter cells autonomously are those
containing cell penetrating peptides (CPPs) (Frankel and Pabo 1988; Green and
Loewenstein 1988; Joliot et al. 1991; Thorén et al. 2000). CPPs have highly
divergent sequences although many are amphipathic or cationic in nature (Drin
et al. 2003). CPPs can mediate uptake of attached cargoes, including low molecular
weight drugs, proteins, nucleic acids and peptides (Rothbard et al. 2000; Nagahara
et al. 1998; Simmons et al. 1997; Shibagaki and Udey 2002; Chugh and Eudes
2008). CPPs may be internalised by virtue of their secondary structure and net
positive charge (Drin et al. 2001, 2003) and there is evidence that their
internalisation can occur via different mechanisms including direct membrane
penetration through pore or inverted micelle formation, clathrin-dependent endocytosis, lipid raft-mediated macropinocytosis or clathrin- and caveolin-independent
endocytosis (Drin et al. 2003; Richard et al. 2005; Wadia et al. 2004; Ter-Avetisyan
et al. 2009; Joliot and Prochiantz 2004). Like CPPs, the uptake domains of AvrM
and AvrL567 have highly divergent sequences and it remains possible that uptake
of these and other fungal effector proteins could depend on the secondary structure
of the N-terminal motifs.
9 Summary
Micro-organisms have adapted to become efficient pathogens of plants,
establishing growth even within living host cells through specialised infection
structures such as haustoria and intracellular invasive hyphae. In biotrophic
interactions where pathogens obtain nutrients from living hosts, pathogens secrete
small proteins known as effectors, which contribute to the establishment and
Plant Infection by Biotrophic Fungal and Oomycete Pathogens
203
maintenance of infection. While considerable progress has been made in
identifying candidate effectors through high throughput screening of pathogen
genome sequences, the functions of these proteins are only just beginning to be
characterised. It is becoming clear that effectors have roles not only in the apoplast
but also within host cells. Key areas for future research include elucidating the
functions of effectors and characterising the mechanism of host intracellular fungal
and oomycete effector delivery into host cells.
References
Agrios GN (1997) Plant pathology. Academic, San Diego
Aist JR (1976) Papillae and related wound plugs of plant cells. Annu Rev Phytopathol 14:145–163
Aist JR, Bushnell WR (1991) Invasion of plants by powdery mildew fungi and cellular
mechanisms of resistance. In: Cole GT, Hoch HC (eds) The fungal spore and disease initiation
in plants and animals. Plenum, New York, pp 321–345
Alfano JR, Collmer A (2004) Type III secretion system effector proteins: double agents in
bacterial disease and plant defense. Annu Rev Phytopathol 42:385–414
An Q, H€uckelhoven R, Kogel KH, van Bel AJE (2006) Multivesicular bodies participate in a cell
wall associated defense response in barley leaves attacked by the pathogenic powdery mildew
fungus. Cell Microbiol 8:1009–1019
Anderson PA, Lawrence GJ, Morrish BC, Ayliffe MA, Finnegan EJ, Ellis JG (1997) Inactivation
of the flax rust resistance gene M associated with loss of a repeated unit within the leucine-rich
repeat coding region. Plant Cell 9:641–651
Aniento F, Robinson DG (2005) Testing for endocytosis in plants. Protoplasma 226:3–11
Avrova AO, Boevink PC, Young V, Grenville-Briggs LJ, Van West P, Birch PRJ, Whisson SC
(2008) A novel Phytophthora infestans haustorium-specific membrane protein is required for
infection of potato. Cell Microbiol 10:2271–2284
Baka ZAM (2008) Occurrence and ultrastructure of Albugo candida on a new host, Arabis alpina
in Saudi Arabia. Micron 39:1138–1144
Baka ZA, Larous L, L€
osel DM (1995) Distribution of ATPase activity at the host-pathogen
interfaces of rust infections. Physiol Mol Plant Pathol 47:67–82
Baldauf SL, Roger AJ, Wenk-Siefert I, Doolittle WF (2000) A kingdom-level phylogeny of
eukaryotes based on combined protein data. Science 290:972–977
Barrett LG, Thrall PH, Dodds PN, van der Merwe M, Linde CC, Lawrence GJ, Burdon JJ (2009)
Diversity and evolution of effector loci in natural populations of the plant pathogen
Melampsora lini. Mol Biol Evol 26:2499–2513
Barth M, Holstein SEH (2004) Identification and functional characterization of Arabidopsis
AP180, a binding partner of plant aC-adaptin. J Cell Sci 117:2051–2062
Bechinger C, Giebel KF, Schnell M, Leiderer P, Deising HB, Bastmeyer M (1999) Optical
measurements of invasive forces exerted by appressoria of a plant pathogenic fungus. Science
285:1896–1899
Birch PRJ, Rehmany AP, Pritchard L, Kamoun S, Beynon JL (2006) Trafficking arms: oomycete
effectors enter host plant cells. Trends Microbiol 14:8–11
Boddey JA, Hodder AN, Gunther S, Gilson PR, Patsiouras H, Kapp EA, Pearce JA, de KoningWard TF, Simpson RJ, Crabb BS, Cowman AF (2010) An aspartyl protease directs malaria
effector proteins to the host cell. Nature 463:627–631
Boller T, Felix G (2009) A renaissance of elicitors: perception of microbe-associated molecular
patterns and danger signals by pattern-recognition receptors. Annu Rev Plant Biol 60:379–406
204
P.H.P. Gan et al.
Borner GHH, Sherrier DJ, Weimar T, Michaelson LV, Hawkins ND, MacAskill A, Napier JA,
Beale MH, Lilley KS, Dupree P (2005) Analysis of detergent-resistant membranes in
Arabidopsis. Evidence for plasma membrane lipid rafts. Plant Physiol 137:104–116
Bos JIB, Kanneganti TD, Young C, Cakir C, Huitema E, Win J, Armstrong MR, Birch PRJ,
Kamoun S (2006) The C-terminal half of Phytophthora infestans RXLR effector AVR3a is
sufficient to trigger R3a-mediated hypersensitivity and suppress INF1-induced cell death in
Nicotiana benthamiana. Plant J 48:165–176
Bos JIB, Chaparro-Garcia A, Quesada-Ocampo LM, Gardener BBM, Kamoun S (2009) Distinct
amino acids of the Phytophthora infestans effector AVR3a condition activation of R3a
hypersensitivity and suppression of cell death. Mol Plant Microbe Interact 22:269–281
Braun EJ, Howard RJ (1994) Adhesion of fungal spores and germlings to host plant surfaces.
Protoplasma 181:202–212
Brunner F, Rosahl S, Lee J, Rudd JJ, Geiler C, Kauppinen S, Rasmussen G, Scheel D, Nurnberger
T (2002) Pep-13, a plant defense-inducing pathogen-associated pattern from Phytophthora
transglutaminases. EMBO J 21:6681–6688
Carver TLW, Thomas BJ, Ingerson-Morris SM (1995) The surface of Erysiphe graminis and the
production of extracellular material at the fungus-host interface during germling and colony
development. Can J Bot 73:272–287
Catanzariti A-M, Dodds PN, Lawrence GJ, Ayliffe MA, Ellis JG (2006) Haustorially expressed
secreted proteins from flax rust are highly enriched for avirulence elicitors. Plant Cell
18:243–256
Catanzariti A-M, Dodds PN, Ve T, Kobe B, Ellis JG, Staskawicz BJ (2010) The AvrM effector
from flax rust has a structured C-terminal domain and interacts directly with the M resistance
protein. Mol Plant Microbe Interact 23:49–57
Champouret N, Bouwmeester K, Rietman H, van der Lee T, Maliepaard C, Heupink A, van de
Vondervoort PJI, Jacobsen E, Visser RGF, van der Vossen EAG, Govers F, Vleeshouwers
VGAA (2009) Phytophthora infestans isolates lacking class I ipiO variants are virulent on
Rpi-blb1 potato. Mol Plant Microbe Interact 22:1535–1545
Choi J, Park J, Kim D, Jung K, Kang S, Lee Y-H (2010) Fungal secretome database: integrated
platform for annotation of fungal secretomes. BMC Genomics 11:105–121
Chugh A, Eudes F (2008) Cellular uptake of cell-penetrating peptides pVEC and transportan in
plants. J Pept Sci 14:477–481
Clement JA, Martin SG, Porter R, Butt TM, Beckett A (1993) Germination and the role of
extracellular matrix in adhesion of urediniospores of Uromyces viciae-fabae to synthetic
surfaces. Mycol Res 97:585–593
Coffey MD (1972) Ultrastructural changes in rust-infected tissues of flax and sunflower. Can J Bot
50:1485–1492
Cornelis GR (2006) The Type III secretion injectisome. Nat Rev Microbiol 4:811–825
Cosio EG, Feger M, Miller CJ, Antelo L, Ebel J (1996) High-affinity binding of fungal b-glucan
elicitors to cell membranes of species of the plant family Fabaceae. Planta 200:92–99
Damasceno CMB, Bishop JG, Ripoll DR, Win J, Kamoun S, Rose JKC (2008) Structure of the
glucanase inhibitor protein (GIP) family from Phytophthora species suggests coevolution with
plant endo-beta-1,3-glucanases. Mol Plant Microbe Interact 21:820–830
de Jonge R, Peter van Esse H, Kombrink A, Shinya T, Desaki Y, Bours R, van der Krol S, Shibuya N,
Joosten MHAJ, Thomma BPHJ (2010) Conserved fungal LysM effector Ecp6 prevents chitintriggered immunity in plants. Science 329:953–955
de Koning-Ward TF, Gilson PR, Boddey JA, Rug M, Smith BJ, Papenfuss AT, Sanders PR, Lundie RJ,
Maier AG, Cowman AF, Crabb BS (2009) A newly discovered protein export machine in malaria
parasites. Nature 459:945–950
Dean RA, Talbot NJ, Ebbole DJ, Farman ML, Mitchell TK, Orbach MJ, Thon M, Kulkarni R, Xu JR,
Pan H (2005) The genome sequence of the rice blast fungus Magnaporthe grisea. Nature
434:980–986
Plant Infection by Biotrophic Fungal and Oomycete Pathogens
205
Deising H, Heiler S, Rauscher M, Xu H, Mendgen K (1996) Cellular aspects of rust infection
structure differentiation. In: Nicole M, Gianinazzi-Pearson V (eds) Histology, ultrastructure
and molecular cytology of plant-microorganism interactions. Kluwer, Dordrecht, pp 135–156
Dhonukshe P, Aniento F, Hwang I, Robinson DG, Mravec J, Stierhof Y-D, Friml J (2007)
Clathrin-mediated constitutive endocytosis of PIN auxin efflux carriers in Arabidopsis. Curr
Biol 17:520–527
Dickinson CH, Lucas JA (1982) Plant pathology and plant pathogens. Blackwell Scientific
Publications, London
Dodds PN, Rathjen JP (2010) Plant immunity: towards an integrated view of plant-pathogen
interactions. Nat Rev Genet 11:539–548
Dodds PN, Lawrence GJ, Catanzariti A-M, Ayliffe MA, Ellis JG (2004) The Melampsora lini
AvrL567 avirulence genes are expressed in haustoria and their products are recognized inside
plant cells. Plant Cell 16:755–768
Dodds PN, Lawrence GJ, Catanzariti A-M, Teh T, Wang C-IA, Ayliffe MA, Kobe B, Ellis JG
(2006) Direct protein interaction underlies gene-for-gene specificity and coevolution of the flax
resistance genes and flax rust avirulence genes. Proc Natl Acad Sci USA 103:8888–8893
D€
oehlemann G (2011) Effectors in Ustilago maydis. In: Perotto S, Baluška F (eds) Signaling and
communication in plant symbiosis. Springer, Heidelberg
Doherty GJ, McMahon HT (2009) Mechanisms of endocytosis. Annu Rev Biochem 78:857–902
Dou D, Kale SD, Wang X, Chen Y, Wang Q, Wang X, Jiang RHY, Arrendondo FD, Anderson RG,
Thakur PB, McDowell JM, Wang Y, Tyler BM (2008a) Conserved C-terminal motifs required
for avirulence and suppresion of cell death by Phytophthora sojae effector Avr1b. Plant Cell
20:1118–1133
Dou D, Kale SD, Wang X, Jiang RHY, Bruce NA, Arredondo FD, Zhang X, Tyler BM (2008b)
RXLR-mediated entry of Phytophthora sojae effector Avr1b into soybean cells does not
require pathogen-encoded machinery. Plant Cell 20:1930–1947
Drin G, Mazel M, Clair P, Mathieu D, Kaczorek M, Temsamani J (2001) Physico-chemical
requirements for cellular uptake of pAntp peptide. Eur J Biochem 268:1304–1314
Drin G, Cottin S, Blanc E, Rees AR, Temsamani J (2003) Studies on the internalization mechanism of cationic cell-penetrating peptides. J Biol Chem 278:31192–31201
Enkerli K, Hahn MG, Mims CW (1997) Ultrastructure of compatible and incompatible
interactions of soybean roots infected with the plant pathogenic oomycete Phytophthora
sojae. Can J Bot 75:1493–1508
Fernandez MR, Heath MC (1991) Interactions of the non-host French bean plant (Phaseolus
vulgaris) parasitic and saprophytic fungi. IV. Effect of preinoculation with the bean rust fungus
on growth of parastic fungi nonpathogenic on beans. Can J Bot 69:1642–1646
Flor HH (1955) Host-parasite interaction in flax rust-its genetics and other implications. Phytopathology 45:680–685
Frankel AD, Pabo CO (1988) Cellular uptake of the Tat protein from the human immunodeficiency
virus. Cell 55:1189–1193
Freytag S, Arabatzis N, Hahlbrock K, Schmelzer E (1994) Reversible cytoplasmic rearrangements
precede wall apposition, hypersensitive cell death and defense-related gene activation in
potato/Phytophthora infestans interactions. Planta 194:123–135
Galan JE, Wolf-Watz H (2006) Protein delivery into eukaryotic cells by type III secretion
machines. Nature 444:567–573
Gan PHP, Rafiqi M, Hardham AR, Dodds PN (2010) Effectors of biotrophic fungal plant
pathogens. Funct Plant Biol 37:913–918
Gil F, Gay JL (1977) Ultrastructural and physiological properties of the host interfacial
components of haustoria of Erysiphe pisi in vivo and in vitro. Physiol Plant Pathol 10:1–12
Godfrey D, Bohlenius H, Pedersen C, Zhang Z, Emmersen J, Thordal-Christensen H (2010)
Powdery mildew and rust fungal effector candidates share N-terminal Y/F/WxC-motif. BMC
Genomics 11:317
206
P.H.P. Gan et al.
Goldberg DE, Cowman AF (2010) Moving in and renovating: exporting proteins from Plasmodium into host erythrocytes. Nat Rev Micro 8:617–621
Gout L, Fudal I, Kuhn ML, Blaise FE, Cattolico L, Balesdent MH, Rouxel T (2006) Lost in the
middle of nowhere: the AvrLm1 avirulence gene of the Dothideomycete Leptosphaeria
maculans. Mol Microbiol 60:67–80
Green M, Loewenstein PM (1988) Autonomous functional domains of chemically synthesized
human immunodeficiency virus tat trans-activator protein. Cell 55:1179–1188
Green JR, Carver TLW, Gurr SJ (2002) The formation and function of infection and feeding
structures. In: Bélanger RR, Bushnell WR, Dik AJ, Carver TLW (eds) Powdery mildews: a
comprehensive treatise. American Phytological Society Press, St Paul, USA, pp 68–82
Grenville-Briggs LJ, Avrova AO, Hay RJ, Bruce CR, Whisson SC, van West P (2010) Identification of appressorial and mycelial cell wall proteins and a survey of the membrane proteome of
Phytophthora infestans. Fungal Biol 114:702–723
Gus-Mayer S, Naton B, Hahlbrock K, Schmelzer E (1998) Local mechanical stimulation induces
components of the pathogen defense response in parsley. Proc Natl Acad Sci USA
95:8398–8403
Haas BJ, Kamoun S, Zody MC, Jiang RHY, Handsaker RE, Cano LM, Grabherr M, Kodira CD,
Raffaele S, Torto-Alalibo T, Bozkurt TO, Ah-Fong AMV, Alvarado L, Anderson VL,
Armstrong MR, Avrova A, Baxter L, Beynon J, Boevink PC, Bollmann SR, Bos JIB, Bulone
V, Cai G, Cakir C, Carrington JC, Chawner M, Conti L, Costanzo S, Ewan R, Fahlgren N,
Fischbach MA, Fugelstad J, Gilroy EM, Gnerre S, Green PJ, Grenville-Briggs LJ, Griffith J,
Grunwald NJ, Horn K, Horner NR, Hu C-H, Huitema E, Jeong D-H, Jones AME, Jones JDG,
Jones RW, Karlsson EK, Kunjeti SG, Lamour K, Liu Z, Ma L, MacLean D, Chibucos MC,
McDonald H, McWalters J, Meijer HJG, Morgan W, Morris PF, Munro CA, O’Neill K,
Ospina-Giraldo M, Pinzon A, Pritchard L, Ramsahoye B, Ren Q, Restrepo S, Roy S,
Sadanandom A, Savidor A, Schornack S, Schwartz DC, Schumann UD, Schwessinger B,
Seyer L, Sharpe T, Silvar C, Song J, Studholme DJ, Sykes S, Thines M, van de Vondervoort
PJI, Phuntumart V, Wawra S, Weide R, Win J, Young C, Zhou S, Fry W, Meyers BC, van West
P, Ristaino J, Govers F, Birch PRJ, Whisson SC, Judelson HS, Nusbaum C (2009) Genome
sequence and analysis of the Irish potato famine pathogen Phytophthora infestans. Nature
461:393–398
Hahn M, Mendgen K (1992) Isolation by ConA binding of haustoria from different rust fungi and
comparison of their surface qualities. Protoplasma 170:95–103
Hahn M, Mendgen K (1997) Characterization of in planta-induced rust genes isolated from a
haustorium-specific cDNA library. Mol Plant Microbe Interact 10:427–437
Hahn M, Deising H, Struck C, Mendgen K (1997) Fungal morphogenesis and enzyme secretion
during pathogenesis. In: Hartleb H, Heitefuss R, Hoppe H-H (eds) Resistance of crop plants
against fungi. Fischer, Jena, pp 33–57
Halterman DA, Chen Y, Sopee J, Berduo-Sandoval J, Sanchez-Perez A (2010) Competition
between Phytophthora infestans effectors leads to increased aggressiveness on plants
containing broad-spectrum late blight resistance. PLoS One 5:e10536
Hamer JE, Howard RJ, Chumley FG, Valent B (1988) A mechanism for surface attachment in
spores of a plant pathogenic fungus. Science 239:288–290
Haney CH, Long SR (2010) Plant flotillins are required for infection by nitrogen-fixing bacteria.
Proc Natl Acad Sci USA 107:478–483
Harder DE, Chong J (1991) Rust haustoria. In: Mendgen K, Lesemann D-E (eds) Electron
microscopy of plant pathogens. Springer, Heidelberg, pp 235–250
Hardham AR (2005) Phytophthora cinnamomi. Mol Plant Pathol 6:589–604
Hardham AR (2007) Cell biology of fungal and Oomycete infection of plants. In: Howard RJ,
Gow NAR (eds) The mycota VII. Springer, Heidelberg, pp 251–289
Hardham AR, Cahill DM (2010) The role of oomycete effectors in plant-pathogen interactions.
Funct Plant Biol 37:919–925
Plant Infection by Biotrophic Fungal and Oomycete Pathogens
207
Hardham A, Takemoto D, White R (2008) Rapid and dynamic subcellular reorganization following mechanical stimulation of Arabidopsis epidermal cells mimics responses to fungal and
oomycete attack. BMC Plant Biol 8:63
Heath MC (1976) Ultrastructural and functional similarity of the haustorial neckband of rust fungi
and the Casparian strip of vascular plants. Can J Bot 54:2482–2489
Heath MC, Skalamera D (1997) Cellular interactions between plants and biotrophic fungal
parasites. Adv Bot Res 24:195–225
Heath MC, Valent B, Howard R, Chumley FG (1991) Interactions of two strains of Magnaporthe
grisea with rice, goosegrass, and weeping lovegrass. Can J Bot 68:1627–1637
Heilmann I (2009) Using genetic tools to understand plant phosphoinositide signalling. Trends
Plant Sci 14:171–179
Hein I, Gilroy EM, Armstrong MR, Birch PRJ (2009) The zig-zag-zig in oomycete–plant
interactions. Mol Plant Pathol 10:547–562
Hoch HC, Staples RC (1991) Signaling for infection structure formation in fungi. In: Cole GT,
Hoch HC (eds) The fungal spore and disease initiation in plants and animals. Plenum, New
York, pp 25–46
Hoch HC, Staples RC, Whitehead B, Comeau J, Wolf ED (1987) Signaling for growth orientation
and cell differentiation by surface topography in Uromyces. Science 235:1659–1662
Holstein SEH, Oliviusson P (2005) Sequence analysis of Arabidopsis thaliana E/ANTH-domaincontaining proteins: membrane tethers of the clathrin-dependent vesicle budding machinery.
Protoplasma 226:13–21
Houterman PM, Cornelissen BJC, Rep M (2008) Suppression of plant resistance gene-based
immunity by a fungal effector. PLoS Pathog 4:e1000061
Howard RJ, Ferrari MA, Roach DH, Money NP (1991) Penetration of hard substrates by a fungus
employing enormous turgor pressures. Proc Natl Acad Sci USA 88:11281–11284
Isberg RR, Tran Van Nhieu G (1994) Binding and internalization of microorganisms by integrin
receptors. Trends Microbiol 2:10–14
Jia Y, McAdams SA, Bryan GT, Hershey HP, Valent B (2000) Direct interaction of resistance
gene and avirulence gene products confers rice blast resistance. EMBO J 19:4004–4014
Jiang RHY, Tripathy S, Govers F, Tyler BM (2008) RXLR effector reservoir in two Phytophthora
species is dominated by a single rapidly evolving superfamily with more than 700 members.
Proc Natl Acad Sci USA 105:4874–4879
Joliot A, Pernelle C, Deagostini-Bazin H, Prochiantz A (1991) Antennapedia homeobox peptide
regulates neural morphogenesis. Proc Natl Acad Sci USA 88:1864–1868
Joly DL, Feau N, Tanguay P, Hamelin RC (2010) Comparative analysis of secreted protein
evolution using expressed sequence tags from four poplar leaf rusts (Melampsora spp.).
BMC Genomics 11:422
Jones JDG, Dangl JL (2006) The plant immune system. Nature 444:323–329
Kaku H, Nishizawa Y, Ishii-Minami N, Akimoto-Tomiyama C, Dohmae N, Takio K, Minami E,
Shibuya N (2006) Plant cells recognize chitin fragments for defense signaling through a plasma
membrane receptor. Proc Natl Acad Sci USA 103:11086–11091
Kale SD, Gu B, Capelluto DGS, Dou D, Feldman E, Rumore A, Arredondo FD, Hanlon R, Fudal I,
Rouxel T, Lawrence CB, Shan W, Tyler BM (2010) External lipid PI3P mediates entry of
eukaryotic pathogen effectors into plant and animal host cells. Cell 142:284–295
Kamper J, Kahmann R, Bolker M, Ma L-J, Brefort T, Saville BJ, Banuett F, Kronstad JW, Gold
SE, Muller O, Perlin MH, Wosten HAB, de Vries R, Ruiz-Herrera J, Reynaga-Pena CG,
Snetselaar K, McCann M, Perez-Martin J, Feldbrugge M, Basse CW, Steinberg G, Ibeas JI,
Holloman W, Guzman P, Farman M, Stajich JE, Sentandreu R, Gonzalez-Prieto JM, Kennell
JC, Molina L, Schirawski J, Mendoza-Mendoza A, Greilinger D, Munch K, Rossel N, Scherer
M, Vranes M, Ladendorf O, Vincon V, Fuchs U, Sandrock B, Meng S, Ho ECH, Cahill MJ,
Boyce KJ, Klose J, Klosterman SJ, Deelstra HJ, Ortiz-Castellanos L, Li W, Sanchez-Alonso P,
Schreier PH, Hauser-Hahn I, Vaupel M, Koopmann E, Friedrich G, Voss H, Schluter T,
Margolis J, Platt D, Swimmer C, Gnirke A, Chen F, Vysotskaia V, Mannhaupt G, Guldener
208
P.H.P. Gan et al.
U, Munsterkotter M, Haase D, Oesterheld M, Mewes H-W, Mauceli EW, DeCaprio D, Wade
CM, Butler J, Young S, Jaffe DB, Calvo S, Nusbaum C, Galagan J, Birren BW (2006) Insights
from the genome of the biotrophic fungal plant pathogen Ustilago maydis. Nature 444:97–101
Kamoun S (2006) A catalogue of the effector secretome of plant pathogenic oomycetes. Annu Rev
Phytopathol 44:41–60
Kankanala P, Czymmek K, Valent B (2007) Roles for rice membrane dynamics and
plasmodesmata during biotrophic invasion by the blast fungus. Plant Cell 19:706–724
Kemen E, Kemen AC, Rafiqi M, Hempel U, Mendgen K, Hahn M, Voegele RT (2005) Identification of a protein from rust fungi transferred from haustoria into infected plant cells. Mol Plant
Microbe Interact 18:1130–1139
Khang CH, Berruyer R, Giraldo MC, Kankanala P, Park S-Y, Czymmek K, Kang S, Valent B
(2010) Translocation of Magnaporthe oryzae effectors into rice cells and their subsequent cellto-cell movement. Plant Cell 22:1388–1403
Kishimoto K, Kouzai Y, Kaku H, Shibuya N, Minami E, Nishizawa Y (2010) Perception of the
chitin oligosaccharides contributes to disease resistance to blast fungus Magnaporthe oryzae in
rice. Plant J 64:343–354
Koh S, André A, Edwards H, Ehrhardt D, Somerville S (2005) Arabidopsis thaliana subcellular
responses to compatible Erysiphe cichoracearum infections. Plant J 44:516–529
Latijnhouwers M, de Wit PJGM, Govers F (2003) Oomycetes and fungi: similar weaponry to
attack plants. Trends Microbiol 11:462–469
Lawrence GJ, Finnegan EJ, Ayliffe MA, Ellis JG (1995) The L6 gene for flax rust resistance is
related to the Arabidopsis bacterial resistance gene RPS2 and the tobacco viral resistance gene
N. Plant Cell 7:1195–1206
Lawrence GJ, Dodds PN, Ellis JG (2010) Transformation of the flax rust fungus, Melampsora lini:
selection via silencing of an avirulence gene. Plant J 61:364–369
Leborgne-Castel N, Adam T, Bouhidel K (2010) Endocytosis in plant-microbe interactions.
Protoplasma 247:177–193
Leckie CP, Callow JA, Green JR (1995) Reorganization of the endoplasmic reticulum in pea leaf
epidermal cells infected by the powdery mildew fungus Erysiphe pisi. New Phytol
131:211–221
Littlefield LJ (1972) Development of haustoria of Melampsora lini. Can J Bot 50:1701–1703
Magzoub M, Pramanik A, Gr€aslund A (2005) Modeling the endosomal escape of cell-penetrating
peptides: transmembrane pH gradient driven translocation across phospholipid bilayers. Biochemistry 44:14890–14897
Manning VA, Ciuffetti LM (2005) Localization of Ptr ToxA produced by Pyrenophora triticirepentis reveals protein import into wheat mesophyll cells. Plant Cell 17:3203–3212
Manning VA, Hamilton SM, Karplus PA, Ciuffetti LM (2008) The Arg-Gly-Asp-containing,
solvent-exposed loop of Ptr ToxA is required for internalization. Mol Plant Microbe Interact
21:315–325
Marcel S, Sawers R, Oakeley E, Angliker H, Paszkowski U (2010) Tissue-adapted invasion
strategies of the rice blast fungus Magnaporthe oryzae. Plant Cell 22:3177–3187
Mayor S, Pagano RE (2007) Pathways of clathrin-independent endocytosis. Nat Rev Mol Cell Biol
8:603–612
Mendgen K, Deising H (1993) Infection structures of fungal plant pathogens – a cytological and
physiological evaluation. New Phytol 124:193–213
Mendgen K, Hahn M (2002) Plant infection and the establishment of fungal biotrophy. Trends
Plant Sci 7:352–356
Mendgen K, Welter K, Scheffold F, Knauf-Bieter G (1991) High pressure freezing of rust infected
leaves. In: Mendgen K, Lesemann D-E (eds) Electron microscopy of plant pathogens.
Springer, Heidelberg, pp 31–42
Mendgen K, Struck C, Voegele RT, Hahn M (2000) Biotrophy and rust haustoria. Physiol Mol
Plant Pathol 56:141–145
Plant Infection by Biotrophic Fungal and Oomycete Pathogens
209
Micali C, Gollner K, Humphry M, Consonni C, Panstruga R (2008) The powdery mildew disease
of Arabidopsis: a paradigm for the interaction between plants and biotrophic fungi. The
Arabidopsis Book
Micali CO, Neumann U, Grunewald D, Panstruga R, O’Connell R (2011) Biogenesis of a
specialized plant-fungal interface during host cell internalization of Golovinomyces orontii
haustoria. Cellular Microbiology 13:210–226
Mims CW, Rodriguez-Lother C, Richardson EA (2002) Ultrastructure of the host-pathogen
interface in daylily leaves infected by the rust fungus Puccinia hemerocallidis. Protoplasma
219:221–226
Mims CW, Celio GJ, Richardson EA (2003) The use of high pressure freezing and freeze
substitution to study host-pathogen interactions in fungal disease of plants. Microsc Microanal
9:522–531
Mithofer A, Fliegmann J, Neuhaus-Url G, Schwarz H, Ebel J (2000) The hepta-b-glucoside
elicitor-binding proteins from legumes represent a putative receptor family. Biol Chem
381:705–713
Mueller O, Kahmann R, Aguilar G, Trejo-Aguilar B, Wu A, de Vries RP (2008) The secretome of
the maize pathogen Ustilago maydis. Fungal Genet Biol 45:S63–S70
M€unch S, Linger U, Floss DS, Liudwig N, Sauer N, Deising H (2008) The hemibiotrophic lifestyle
of Colletotrichum species. J Plant Physiol 165:41–51
Murdoch LJ, Hardham AR (1998) Components in the haustorial wall of the flax rust fungus,
Melampsora lini, are labelled by three anti-calmodulin monoclonal antibodies. Protoplasma
201:180–193
Nagahara H, Vocero-Akbani AM, Snyder EL, Ho A, Latham DG, Lissy NA, Becker-Hapak M,
Ezhevsky SA, Dowdy SF (1998) Transduction of full-length TAT fusion proteins into mammalian cells: TAT-p27Kip1 induces cell migration. Nat Med 4:1449–1452
O’Connell R (1987) Absence of a specialised interface between infection hyphae of
Colletotrichum lindemuthianum and Phaseolus vulgaris. New Phytol 107:725–734
O’Connell RJ, Panstruga R (2006) Tête à tête inside a plant cell: establishing compatibility
between plants and biotrophic fungi and oomycetes. New Phytol 171:699–718
Oh S-K, Young C, Lee M, Oliva R, Bozkurt TO, Cano LM, Win J, Bos JIB, Liu H-Y, van Damme
M, Morgan W, Choi D, Van der Vossen EAG, Vleeshouwers VGAA, Kamoun S (2009)
In planta expression screens of Phytophthora infestans RXLR effectors reveal diverse
phenotypes, including activation of the Solanum bulbocastanum disease resistance protein
Rpi-blb2. Plant Cell 21:2928–2947
Orbach MJ, Farrall L, Sweigard JA, Chumley FG, Valent B (2000) A telomeric avirulence gene
determines efficacy for the rice blast resistance gene Pi-ta. Plant Cell 12:2019–2032
Perfect SE, Green JR (2001) Infection structures of biotrophic and hemibiotrophic fungal plant
pathogens. Mol Plant Pathol 2:101–108
Rafiqi M, Gan P, Ravensdale M, Lawrence GJ, Ellis JG, Jones DA, Hardham AR, Dodds PN
(2010) Internalization of flax rust avirulence proteins into flax and tobacco cells can occur in
the absence of the pathogen. Plant Cell 22:2017–2032
Read ND, Kellock HK, Trewavas AJ (1992) Contact sensing during infection by fungal pathogens.
In: Callow A, Green R (eds) Perspectives in plant cell recognition. Cambridge University
Press, Cambridge, pp 137–172
Rehmany AP, Gordon A, Rose LE, Allen RL, Armstrong MR, Whisson SC, Kamoun S, Tyler BM,
Birch PRJ, Beynon JL (2005) Differential recognition of highly divergent downy mildew
avirulence gene alleles by RPP1 resistance genes from two Arabidopsis lines. Plant Cell
17:1839–1850
Richard JP, Melikov K, Brooks H, Prevot P, Lebleu B, Chernomordik LV (2005) Cellular uptake
of unconjugated TAT peptide involves clathrin-dependent endocytosis and heparan sulfate
receptors. J Biol Chem 280:15300–15306
210
P.H.P. Gan et al.
Ridout CJ, Skamnioti P, Porritt O, Sacristan S, Jones JDG, Brown JKM (2006) Multiple avirulence
paralogues in cereal powdery mildew fungi may contribute to parasite fitness and defeat of
plant resistance. Plant Cell 18:2402–2414
Roberts AM, Mackie AJ, Hathaway V, Callow JA, Green JR (1993) Molecular differentiation in
the extrahaustorial membrane of pea powdery mildew haustoria at early and late stages of
development. Physiol Mol Plant Pathol 43:147–160
Rooney HCE, van’t Klooster JW, van der Hoorn RAL, Joosten MHAJ, Jones JDG, de Wit PJGM
(2005) Cladosporium Avr2 inhibits tomato Rcr3 protease required for Cf-2-dependent disease
resistance. Science 308:1783–1786
Rose JKC, Ham KS, Darvill AG, Albersheim P (2002) Molecular cloning and characterization of
glucanase inhibitor proteins: coevolution of a counterdefense mechanism by plant pathogens.
Plant Cell 14:1329–1345
Rothbard JB, Garlington S, Lin Q, Kirschberg T, Kreider E, McGrane PL, Wender PA, Khavari
PA (2000) Conjugation of arginine oligomers to cyclosporin A facilitates topical delivery and
inhibition of inflammation. Nat Med 6:1253–1257
Sandvig K, van Deurs B (1996) Endocytosis, intracellular transport, and cytotoxic action of shiga
toxin and ricin. Physiol Rev 76:949–966
Sandvig K, van Deurs B (2002) Membrane traffic exploited by protein toxins. Annu Rev Cell Dev
Biol 18:1–24
Scholes J, Farrar J (1985) Photosynthesis and chloroplast functioning within individual pustules of
Uromyces muscari on bluebell leaves. Physiol Plant Pathol 27:387–400
Schornack S, Van Damme M, Bozkurt TO, Cano LM, Smoker M, Thines M, Gaulin E, Kamoun S,
Huitema E (2010) Ancient class of translocated oomycete effectors targets the host nucleus.
Proc Natl Acad Sci USA. 107:17421–17426
Schwessinger B, Zipfel C (2008) News from the frontline: recent insights into PAMP-triggered
immunity in plants. Curr Opin Plant Biol 11:389–395
Senchou V, Weide R, Carrasco A, Bouyssou H, Pont-Lezica R, Govers F, Canut H (2004) High
affinity recognition of a Phytophthora protein by Arabidopsis via an RGD motif. Cell Mol Life
Sci 61:502–509
Sesma A, Osbourn AE (2004) The rice leaf blast pathogen undergoes developmental processes
typical of root-infecting fungi. Nature 431:582–586
Shabab M, Shindo T, Gu C, Kaschani F, Pansuriya T, Chintha R, Harzen A, Colby T, Kamoun S,
van der Hoorn RAL (2008) Fungal effector protein AVR2 targets diversifying defense-related
cys proteases of tomato. Plant Cell 20:1169–1183
Shibagaki N, Udey MC (2002) Dendritic cells transduced with protein antigens induce cytotoxic
lymphocytes and elicit antitumor immunity. J Immunol 168:2393–2401
Simmons CG, Pitts AE, Mayfield LD, Shay JW, Corey DR (1997) Synthesis and membrane
permeability of PNA-peptide conjugates. Bioorg Med Chem Lett 7:3001–3006
Skibbe DS, Doehlemann G, Fernandes J, Walbot V (2010) Maize tumors caused by Ustilago
maydis require organ-specific genes in host and pathogen. Science 328:89–92
Song J, Win J, Tian M, Schornack S, Kaschani F, Ilyas M, van der Hoorn RAL, Kamoun S (2009)
Apoplastic effectors secreted by two unrelated eukaryotic plant pathogens target the tomato
defense protease Rcr3. Proc Natl Acad Sci USA 106:1654–1659
Soylu S (2004) Ultrastructural chracterisation of the host-pathogen interface in white blisterinfected Arabidopsis leaves. Mycopathologia 158:457–464
Staples RC, Hoch HC (1997) Physical and chemical cues for spore germination and appressorium
formation by fungal pathogens. In: Carroll G, Tudzynski P (eds) The mycota V part A.
Springer, Berlin, pp 27–40
Staskawicz BJ, Mudgett MB, Dangl JL, Galan JE (2001) Common and contrasting themes of plant
and animal diseases. Science 292:2285–2289
Struck C, Hahn M, Mendgen K (1996) Plasma membrane H+-ATPase activity in spores, germ
tubes, and haustoria of the rust fungus Uromyces viciae-fabae. Fungal Genet Biol 20:30–35
Plant Infection by Biotrophic Fungal and Oomycete Pathogens
211
Struck C, Siebels C, Rommel O, Wernitz M, Hahn M (1998) The plasma membrane H+-ATPase
from the biotrophic rust fungus Uromyces fabae: molecular characterization of the gene
(PMA1) and functional expression of the enzyme in yeast. Mol Plant Microbe Interact
11:458–465
Struck C, Ernst M, Hahn M (2002) Characterization of a developmentally regulated amino acid
transporter (AAT1p) of the rust fungus Uromyces fabae. Mol Plant Pathol 3:23–30
Szabo LJ, Bushnell WR (2001) Hidden robbers: the role of fungal haustoria in parasitism of plants.
Proc Natl Acad Sci USA 98:7654–7655
Takemoto D, Jones DA, Hardham AR (2003) GFP-tagging of cell components reveals the
dynamics of subcellular re-organization in response to infection of Arabidopsis by oomycete
pathogens. Plant J 33:775–792
Talbot NJ (2003) On the trail of a cereal killer: exploring the biology of Magnaporthe grisea. Annu
Rev Microbiol 57:177–202
Ter-Avetisyan G, T€unnemann G, Nowak D, Nitschke M, Hermann A, Drab M, Cardoso MC
(2009) Cell entry of arginine-rich peptides is independent of endocytosis. J Biol Chem
284:3370–3378
Thorén PEG, Persson D, Karlsson M, Nordén B (2000) The Antennapedia peptide penetratin
translocates across lipid bilayers – the first direct observation. FEBS Lett 482:265–268
Tian M, Benedetti B, Kamoun S (2005) A second Kazal-like protease inhibitor from Phytophthora
infestans inhibits and interacts with the apoplastic pathogenesis-related protease P69B of
tomato. Plant Physiol 138:1785–1793
Tian M, Win J, Song J, van der Hoorn R, van der Knaap E, Kamoun S (2007) A Phytophthora
infestans cystatin-like protein targets a novel tomato papain-like apoplastic protease. Plant
Physiol 143:364–377
Tiburzy R, Martins EMF, Reisener HJ (1992) Isolation of haustoria of Puccinia graminis f. sp.
tritici from wheat leaves. Experiment Mycol 16:324–328
Torres MA (2010) ROS in biotic interactions. Physiol Plant 138:414–429
Torto TA, Li S, Styer A, Huitema E, Testa A, Gow NAR, van West P, Kamoun S (2003) EST
mining and functional expression assays identify extracellular effector proteins from the plant
pathogen Phytophthora. Genome Res 13:1675–1685
Tucker SL, Besi MI, Galhano R, Franceschetti M, Goetz S, Lenhert S, Osbourn A, Sesma A (2010)
Common genetic pathways regulate organ-specific infection-related development in the rice
blast fungus. Plant Cell 22:953–972
Tyler BM, Tripathy S, Zhang X, Dehal P, Jiang RHY, Aerts A, Arredondo FD, Baxter L,
Bensasson D, Beynon JL, Chapman J, Damasceno CMB, Dorrance AE, Dou D, Dickerman
AW, Dubchak IL, Garbelotto M, Gijzen M, Gordon SG, Govers F, Grunwald NJ, Huang W,
Ivors KL, Jones RW, Kamoun S, Krampis K, Lamour KH, Lee M-K, McDonald WH, Medina
M, Meijer HJG, Nordberg EK, Maclean DJ, Ospina-Giraldo MD, Morris PF, Phuntumart V,
Putnam NH, Rash S, Rose JKC, Sakihama Y, Salamov AA, Savidor A, Scheuring CF, Smith
BM, Sobral BWS, Terry A, Torto-Alalibo TA, Win J, Xu Z, Zhang H, Grigoriev IV, Rokhsar
DS, Boore JL (2006) Phytophthora genome sequences uncover evolutionary origins and
mechanisms of pathogenesis. Science 313:1261–1266
Van de Peer Y, De Watcher R (1997) Evolutionary relationships among the eukaryotic crown taxa
taking into account site-to-site rate variation in 18S rRNA. J Mol Evol 45:619–630
van den Burg HA, Harrison SJ, Joosten MHAJ, Vervoort J, de Wit PJGM (2006) Cladosporium
fulvum Avr4 protects fungal cell walls against hydrolysis by plant chitinases accumulating
during infection. Mol Plant Microbe Interact 19:1420–1430
van Esse HP, van’t Klooster JW, Bolton MD, Yadeta KA, van Baarlen P, Boeren S, Vervoort J, de
Wit PJGM, Thomma BPHJ (2008) The Cladosporium fulvum virulence protein Avr2 inhibits
host proteases required for basal defense. Plant Cell 20:1948–1963
van West P, de Bruijn I, Minor KL, Phillips AJ, Robertson EJ, Wawra S, Bain J, Anderson VL,
Secombes CJ (2010) The putative RxLR effector protein SpHtp1 from the fish pathogenic
212
P.H.P. Gan et al.
oomycete Saprolegnia parasitica is translocated into fish cells. FEMS Microbiol Lett
310:127–137
Voegele RT (2006) Uromyces fabae: development, metabolism, and interactions with its host
Vicia faba. FEMS Microbiol Lett 259:165–173
Voegele RT, Mendgen K (2003) Rust haustoria: nutrient uptake and beyond. New Phytol
159:93–100
Voegele RT, Struck C, Hahn M, Mendgen K (2001) The role of haustoria in sugar supply during
infection of broad bean by the rust fungus Uromyces fabae. Proc Natl Acad Sci USA
98:8133–8138
Voegele RT, Hahn M, Mendgen K (2009) The Uredinales: cytology, biochemistry, and molecular
biology. In: Esser K (ed) The mycota. Springer, Berlin, pp 69–98
Wadia JS, Stan RV, Dowdy SF (2004) Transducible TAT-HA fusogenic peptide enhances escape
of TAT-fusion proteins after lipid raft macropinocytosis. Nat Med 10:310–315
Whisson SC, Boevink PC, Moleleki L, Avrova AO, Morales JG, Gilroy EM, Armstrong MR,
Grouffaud S, van West P, Chapman S, Hein I, Toth IK, Pritchard L, Birch PRJ (2007)
A translocation signal for delivery of oomycete effector proteins into host plant cells. Nature
450:115–118
Willmer C, Fricker M (1996) Stomata. Chapman and Hall, Suffolk, Great Britain
Wilson RA, Talbot NJ (2009) Under pressure: investigating the biology of plant infection by
Magnaporthe oryzae. Nat Rev Microbiol 7:185–195
Win J, Morgan W, Bos J, Krasileva KV, Cano LM, Chaparro-Garcia A, Ammar R, Staskawicz BJ,
Kamoun S (2007) Adaptive evolution has targeted the C-terminal domain of the RXLR
effectors of plant pathogenic oomycetes. Plant Cell 19:2349–2369
Xu H, Mendgen K (1994) Endocytosis of 1,3-b-glucans by broad bean cells at the penetration site
of the cowpea rust fungus (haploid stage). Planta 195:282–290
Compatibility in Biotrophic Plant–Fungal
Interactions: Ustilago maydis and Friends
Kerstin Schipper and Gunther Doehlemann
Abstract Biotrophic plant pathogens depend on living host cells during all stages
of pathogenic interaction. Ustilago maydis, the causative agent of smut disease
induces development of tumors on all aerial organs of its host plant maize. Immediately upon host penetration, biotrophy is established and maintained during
fungal proliferation and nutrition up to the formation of sexual spores. This requires
an efficient suppression of plant defense responses, in particular host cell death. In
the molecular communication between pathogen and its host, secreted effector
proteins play essential roles. The actual functions of these effectors, however,
still remain largely elusive. To successfully execute the different steps of pathogenic interaction, a tight regulatory network has evolved in the pathogens,
coordinating expression of secreted effectors in a stage- and organ-specific manner.
In this chapter, we discuss the complex molecular mechanisms that ensure compatibility in the intimate relationship between biotrophic fungi and their plant hosts.
1 Introduction
During evolution biotrophic fungi developed a multiplicity of strategies to cope
with their plant hosts. Mechanisms to establish compatible biotrophic interactions
mainly involve the escape from host recognition as well as manipulation of the
plant defense systems and reprogramming of its metabolism. In this chapter, we
will highlight the key features of fungal biotrophic plant pathogens. The basidiomycete smut fungus Ustilago maydis that parasitizes on corn represents an important model organism for biotrophic plant–fungal interactions and will be discussed
K. Schipper • G. Doehlemann (*)
Max Planck Institute for Terrestrial Microbiology, Karl-von-Frisch-Straße 10, 35043 Marburg,
Germany
e-mail: [email protected]
S. Perotto and F. Baluska (eds.), Signaling and Communication in Plant Symbiosis,
Signaling and Communication in Plants 11, DOI 10.1007/978-3-642-20966-6_9,
# Springer-Verlag Berlin Heidelberg 2012
213
214
K. Schipper and G. Doehlemann
in detail. Particular emphasis will lie on the crucial role of secreted effectors in the
crosstalk between the parasite and its host.
1.1
Definition of Biotrophic Plant–Parasite Interactions
One way to classify plant pathogens is according to their feeding strategy (Oliver
and Ipcho 2004). By definition biotrophic plant parasites rely on living substrates.
Many of them establish long-term feeding relationships with their hosts (Lewis
1973; Staples 2000; Mendgen and Hahn 2002). These characteristics are also
resembled by the term biotrophy that has been derived from the Greek words bios
(“life”) and trophy (“feeding”). In contrast to biotrophs, necrotrophic pathogens
(e.g., Sclerotina sclerotioum, Botrytis cinerea) developed a feeding strategy based
on destroying of the host tissue to obtain food from the dead plant material (Lewis
1973). Therefore, necrotrophs kill host cells by secreted toxins which allow the
parasites to grow saprophytically on the released nutrients (Farrar 1984; Walton
1996; van Kan 2006).
To succeed in their intimate host relationship, biotrophs have evolved strategies
to keep the invaded plant cells alive during infection. The entire pathogenic
development including adhesion, penetration, host colonization as well as the
completion of the life cycle by spore formation needs to occur with minimal
damage of the host. Major challenges of biotrophic pathogens to assure compatibility are therefore on one hand to mask themselves in order to circumvent
recognition by the host and on the other hand, to suppress those host defense
mechanisms that cannot be avoided (van Esse et al. 2007; de Wit 2007). Parasitic
biotrophs obtain nutrients from shoot tissue, having no alternative energy sources.
Thus, the host metabolism needs to be reprogrammed in a way that ensures a
continuous supply of nutrients to the pathogen (Spanu and K€amper 2010).
Biotrophy is not restricted to a single taxonomic group of plant pathogens but
exists in different kingdoms ranging from fungi (i.e., basidiomycetes like rusts and
smuts; ascomycetes like powdery mildews) and oomycetes (i.e., downy mildews) to
the bacteria (i.e., Pseudomonas syringae, Xanthomonas campestris). Even the
animal kingdom is represented by plant-parasitizing root-knot and cyst nematodes.
It is not yet clear whether biotrophy is a result of convergent evolution in the
different taxonomic groups or if this lifestyle has a very early common evolutionary
origin (Schulze-Lefert and Panstruga 2003). An interesting view on biotrophy was
given by the finding that endophytic fungi hold the potential being pathogens. This
was nicely shown in recent studies on the Epichloe¨/Neotyphodium endophytes that
usually colonize rye grass in symbiotic interactions (Tanaka et al. 2006, 2008).
In this light, the evolutionary question about emergence of biotrophy might be
asked: are we dealing with a preliminary stage of development towards a mutualistic interaction, or might symbionts represent pathogens that are tamed by their
respective host plants?
Compatibility in Biotrophic Plant–Fungal Interactions: Ustilago maydis and Friends
215
Biotrophic organisms comprise important plant pests that cause enormous economic losses of agricultural plants worldwide. Furthermore, in natural environments they can reduce the competitive abilities of the host. While economically not
threatening biotrophs only lead to moderate agricultural problems, others are much
more harmful pests. Prominent examples of devastating plant pathogenic fungal
biotrophs are the cereal rusts like Puccinia graminis f. sp. tritici (Staples 2003). In
the second half of the nineteenth century, the coffee bean rust Hemileia vastatrix
caused devastating losses in Sri Lanka. This led to collapse of the coffee industry,
which is the reason for this region mainly producing tea and rubber nowadays
(Waller et al. 2007).
Biotrophic host–parasite relationships can be maintained either transiently or
during the complete pathogenic program up to sporulation. Transient types of
biotrophy that are referred to as hemibiotrophy are observed, e.g., in the fungi
Magnaporthe grisea, in different Colletotrichum subspecies, and in the oomycete
pest Phytophthora infestans (Latunde-Dada 2001; Mendgen and Hahn 2002;
Hammond-Kosack and Parker 2003; Haldar et al. 2006). Here, the parasite initially
establishes a biotrophic interaction with its host plant that switches to a
necrotrophic stage while the infection proceeds (Perfect and Green 2001).
Upon the “true” biotrophs that stay biotrophic throughout their complete life
cycle obligate and nonobligate parasites must be differentiated. Obligate biotrophs
are in many cases difficult or not at all cultivable in the absence of their host plant,
which causes immense experimental disadvantages in their study. On the other
hand, nonobligate biotrophs like U. maydis are able to grow under artificial laboratory conditions and are amenable to genetic modifications. Therefore, they often
deal as model systems for biotrophy (Brefort et al. 2009).
2 Pathogenic Development of Biotrophic Fungi
Although being capable of propagating saprophytically, nonobligate biotrophs rely
on the biotrophic phase for sexual recombination. Most biotrophic fungi share a
common infection program. First stages of pathogenic development include host
recognition, attachment to the surface, and penetration at suitable surface sites. The
differentiation of specialized infection structures termed appressoria is widely
spread upon plant-pathogenic fungi. This structure mediates a direct penetration
of the plant tissue, an important stage that resembles the switch from extracellular to
invasive biotrophic growth. Therefore, the penetration event is considered to be a
complex process. Some pathogenic fungi like the rice blast M. grisea or
Colletotrichum spp. facilitate invasion of epidermal cells by mechanical force
(Howard et al. 1991; Money and Howard 1996). In this case, penetration is promoted by an enormous turgor pressure that develops in the dome-shaped, melanized
appressoria of these pathogens (i.e., 8 MPa in M. grisea; Tucker and Talbot 2001).
Alternatively, host penetration may be mediated by the local secretion of lytic
enzymes or by a combination of both mechanisms employing slightly lower turgor
216
K. Schipper and G. Doehlemann
pressures, i.e., as it has been observed in Blumeria graminis (Bailey et al. 1992;
Francis et al. 1996; Pryce-Jones et al. 1999). In the latter cases, infection structures
are usually less prominent and nonmelanized (Mendgen and Deising 1993).
Following penetration, colonization of the plant occurs in the host tissue.
Although biotrophs all rely on living material during plant colonization, the actual
relationship to the host is realized in multiple ways. Spreading in the plant can occur
subcuticular like, e.g., observed in plant infections with the ascomycete Venturia
inaequalis, causing apple scab disease. Some pathogens like U. maydis, Claviceps
purpurea, and monokaryotic rust fungi spread inter- as well as intracellularly.
Powdery mildews, dicaryotic rust fungi, and downy mildews grow mainly extraor intercellular but develop specialized intracellular feeding structures which can be
situated either in epidermal or parenchymal tissue (reviewed in Mendgen and Hahn
2002).
Although plant colonization proceeds in many different ways, a common principle of invasive growth is the invagination of the host plasma membrane, which
creates a matrix-filled interface between the parasite and the host cell, termed the
biotrophic interface (Bauer et al. 1997). This unique compartment is the site of
communication between pathogen and host and thus, probably the most important
scenery during the establishment of the biotrophic interaction. In the past years it
has become more and more evident that proteins secreted to the interface are the
mediators of parasite–host communication. Remarkably, a subset of the weaponry
of secreted effectors the pathogen sends to the interaction zone even enters the plant
cell where the proteins function either in the cytoplasm or in organelles like the
nucleus (see Sect. 3; Kemen et al. 2005; Khang et al. 2010).
An additional characteristic feature of many (e.g., rust fungi, powdery mildews)
but not all biotrophic fungi is the development of intracellular feeding structures,
the so-called haustoria. Typically, haustorium-forming biotrophs grow extra- or
intercellularly, invading only a few plant cells which are then completely filled with
fungal material. Thus, a nutrient sink is generated at the infection site to ensure
feeding of the parasite (Mendgen and Hahn 2002; O’Connell and Panstruga 2006).
The extended haustorial surface area might provide a perfect prerequisite for
nutrient flow towards the parasite. Some biotrophs like U. maydis do not differentiate haustoria (Brefort et al. 2009). Nevertheless, it has been demonstrated that those
pathogens are also able to establish a nutritional sink tissue at the infection site
(Doehlemann et al. 2008b; Horst et al. 2008), probably mediated by the biotrophic
interfaces surrounding intracellular growing hyphae.
3 Ustilago maydis as a Model System for Biotrophic Fungi
U. maydis is the leading model organism of the plant pathogenic smut fungi. The
basidiomycete fungus parasitizes only on Zea mays and its wild progenitor teosinte
(Z. mays ssp. mexicana and ssp. pavigluminis) where it causes corn smut disease.
Concomitantly, U. maydis induces the formation of plant tumors in which the
Compatibility in Biotrophic Plant–Fungal Interactions: Ustilago maydis and Friends
217
fungus completes its disease cycle (Christensen 1963; Banuett 1995; Banuett and
Herskowitz 1996; Doehlemann et al. 2008a).
U. maydis has a diphasic life cycle: the fungus grows saprophytically as haploid
sporidia but cannot cause disease in this form. To infect maize plants, sporidia of
different mating type need to recognize each other on the host surface via a
pheromone–receptor system (B€olker et al. 1992). Perception of the compatible
pheromone induces formation of conjugation tubes that fuse at their tips and thus
form the infectious dicaryotic filament (Sleumer 1932; Rowell 1955; Snetselaar
1993; Snetselaar et al. 1996). This filament is able to enter the plant tissue directly,
which is accompanied by swelling of the hyphal tip resulting in an appressorial
structure (Snetselaar and Mims 1992). Immediately upon host invasion, the
biotrophic relationship is established and fungal hyphae grow intracellularly in
the infected host tissue, being surrounded by a biotrophic interface (Fig. 1).
Biotrophic hyhae proliferate massively, particularly in mesophyl and vascular
tissue until basidiospore formation occurs (Snetselaar and Mims 1994; Doehlemann
et al. 2008a). This goes along with formation of tumors on all areal parts of the
infected maize plants (Brefort et al. 2009). The diploid spores are released when
the tumors dry up and rupture. Under favorable conditions these spores germinate,
the diploid nucleus undergoes meiosis, and budding off from a promycelium,
Fig. 1 Potential sites of U. maydis effector activity. The figure represents a schematic view of a
maize cell that has been invaded by an intracellular growing fungal hypha. During this process, the
plant plasma membrane invaginates and the fungal hypha is separated from the plant symplast by
the biotrophic interface. Since adult maize cells contain a large central vacuole the cytoplasm
containing cell organelles resembles up only a thin layer directly adjacent to the plant cell wall and
the fungal hypha. Secreted effectors localize to the biotrophic interaction zone (open triangles).
Moreover, effectors potentially enter the plant cell (dark triangles). The close up view visualizes
three different putative functions of the secreted apoplastic effectors
218
K. Schipper and G. Doehlemann
haploid sporidia are again produced (Christensen 1963; Snetselaar and Mims 1994;
Banuett 1995; Banuett and Herskowitz 1996; Kahmann et al. 2000).
The U. maydis–Z. mays pathosystem has become more and more important not
only as a model for plant pathogenic smut fungi but also for one of the few models
for “true” biotrophic host–pathogen interactions (K€amper et al. 2006; Brefort et al.
2009). In contrast to obligate biotrophs, haploid U. maydis cells can be efficiently
grown in synthetic media and genetic modifications can be performed with a
convenient toolbox of established methods available, i.e., an efficient gene replacement system, a set of constitutive and inducible promoters as well as genetically
modified strains that grow filamentously in axenic culture (K€amper 2004; Spellig
et al. 1996; Brachmann et al. 2001; Zarnack et al. 2006). In addition, advanced life
cell imaging and a variety of staining methods have been established for U. maydis
to characterize in planta development of the fungus (Basse and Steinberg 2004;
Doehlemann et al. 2008a, b; Brefort et al. 2009).
The 20.5-Mb genome of U. maydis has been sequenced, manually refined and
sequences are publicly available as well as regularly updated on a MIPS platform
(http://www.broad.mit.edu/annotation/genome/ustilago_maydis/Home.html; http://
mips.gsf.de/genre/proj/ustilago/; K€amper et al. 2006). Genome availability furthermore laid the foundation for the establishment of DNA microarray profiling (K€amper
et al. 2006).
For U. maydis infection, sporidia of compatible mating types have to meet and
fuse on the leaf surface, leading to the formation of a diploid infectious filament.
The development of solopathogenic strains that do not need compatible mating
partners has further improved the pathosystem allowing the fast generation
and subsequent phenotyping of U. maydis mutants with no longer need to introduce
mutations in two haploid wild-type strains (B€olker et al. 1995; K€amper et al. 2006).
Disease assays can be performed on young maize seedlings with development
of symptoms (i.e., chlorosis, anthocyan, tumors) within two weeks after
sowing (Banuett 1995). Standardized disease rating combined with confocal microscopic imaging techniques and PCR-based quantification of fungal biomass
permits not only discrimination between infected or noninfected plants but also
allows specification of symptom severity as well as the degree and manner of
intra- and intercellular proliferation of the pathogen (K€amper et al. 2006;
Doehlemann et al. 2008b; di Stasio et al. 2009). Thus, this system allows not only
the identification of essential virulence factors but also of U. maydis mutants with
slightly altered virulence ranging from minor reductions to hypervirulence
(K€amper et al. 2006).
Genetic modifications of the host plant maize like gene silencing or gene
insertions are possible though very time consuming (Shrawat and L€orz 2006).
However, a systemic virus-induced gene silencing system for maize has recently
been developed to study host genes during U. maydis infection. This tool will
increase the methodological potential for future studies on this pathosystem (van
der Linde et al. 2010).
Compatibility in Biotrophic Plant–Fungal Interactions: Ustilago maydis and Friends
219
4 Plant Responses to U. maydis
Plants have evolved multifaceted defense mechanisms against pathogen attacks. In
a first line of defense, recognition of conserved microbial molecules (pathogenassociated molecular patterns, PAMPS) elicits a basal defense response
characterized by the induction of pathogenesis-related (PR) genes, strengthening
of cell walls as well as the production of secondary metabolites. Often, these
responses are associated with the production of reactive oxygen species and the
induction of localized cell death (the hypersensitive response, HR), leading to a
PAMP-triggered resistance (Jones and Dangl 2006).
Specific virulence factors suppress PAMP-triggered responses. However, they again
might be recognized by plant resistance genes that developed in a co-evolutionary arms
race. Defense responses activated by resistance genes involve HR induction,
associated with changes in phytohormone levels. A process leading to the so-called,
effector-triggered immunity (Jones and Dangl 2006), The phytohormone changes
are specifically adapted to the attacking pathogen. The classical, widely accepted
model describes salicylic acid (SA)-dependent defenses acting against biotrophic
pathogens. In contrast, jasmonate (JA)- as well as ethylene-dependent responses
were shown to defend infections with necrotrophs (Greenberg and Yao 2004;
Glazebrook 2005; Jones and Dangl 2006; O’Connell and Panstruga 2006).
In the compatible U. maydis–Z. mays pathosystem indications of a transient
induction of plant defense responses can be recognized on the micro- and macroscopic level, which is changed upon successful establishment of biotrophy about
20–24 h after inoculation (Doehlemann et al. 2008b). During the early phase of plant
colonization, intracellular U. maydis hyphae grow at the apex thereby moving their
cytoplasm to the tip compartment. At the same time older hyphal parts remain empty
and are sealed off by septae (Snetselaar and Mims 1994). Hyphal autofluorescence
develops at the empty sections, indicating enhanced recognition of these old parts by
the plant. Finally, plant cells containing such collapsed hyphae undergo cell death
(Doehlemann et al. 2008b). These processes coincide with early infection symptoms
which are chlorosis and, eventually, small necrotic spots that develop at the infection sites and which are most likely caused by single dying plant cells.
Later on, in colonized tissues, strong autofluorescence and occasional small
areas with clusters of dead cells were detected, indicating enhanced plant defense
sporadically elicited by some hyphae (Doehlemann et al. 2008b). Finally, U.
maydis-induced tumors are formed by enlargement as well as proliferation of
plant cells which goes along with massive proliferation of the fungus. Although
large fungal aggregates are formed in intercellular spaces, programmed cell death
(PCD) can only rarely be observed in the surrounding plant tissue (Doehlemann
et al. 2008a, b).
To investigate plant reactions on the molecular level, transcriptional changes of
maize in response to U. maydis infection were studied by a differential display
approach (Basse 2005) as well as by a time-resolving analysis using the Affymetrix
DNA microarray system (Doehlemann et al. 2008b). Microarray analyses revealed
220
K. Schipper and G. Doehlemann
that an initial recognition of the fungus takes place on the leaf surface, eliciting a
strong but rather unspecific plant defense response. The transient induction of plant
defense genes suggests that U. maydis is recognized via PAMPs. In accordance with
this hypothesis, maize orthologs of specific PAMP-induced LRR-like receptor-like
kinases were shown to be transcriptionally induced during this early phase of
infection (Doehlemann et al. 2008b; Zipfel et al. 2004, 2006). Additionally, a
SER kinase belonging to a group that includes BAK1 was up-regulated
(Doehlemann et al. 2008b). BAK1 has been shown to act as a positive regulator
in infection-induced cell death signaling (Chinchilla et al. 2007; Kemmerling et al.
2007). These observations indicate that PAMP-induced signaling is indeed
activated during the penetration phase of U. maydis resulting in a strong and
unspecific plant response.
However, with establishment of the biotrophic interaction 24 h after infection,
the initial unspecific defense responses were attenuated (Doehlemann et al. 2008b).
Evidence for this was provided by the observation that previously induced defense
gene expression decreased significantly with the onset of biotrophy. In parallel,
induction of genes encoding cell death suppressors such as Bax-Inhibitor-1
(Eichmann et al. 2004) as well as the repression of caspases was observed
(Doehlemann et al. 2008b). These transcriptional changes ensure the secondary
suppression of the initially elicited unspecific defense mechanisms which is a
prerequisite for establishment of biotrophy.
The specific plant response to U. maydis infection is also mirrored by transcriptional changes in plant hormone signaling. During compatible interactions,
biotrophic pathogens induce primarily JA and ethylene responses. These hormones
act antagonistic to SA signaling which counteracts biotrophic pathogens and are
associated with induction of tryptophane biosynthesis, the accumulation of secondary metabolites, and the induction of plant genes encoding defensins (Brader et al.
2001; Glazebrook 2005; Wasternack 2007). In line with this, an induction of JA
signaling as well as typical JA-responsive defense genes such as defensins and
Bowman–Birk-like proteinase inhibitors was also observed during U. maydis
infections (Basse 2005; Doehlemann et al. 2008b). Moreover, transcripts of PR1,
a marker protein for SA-dependent cell death responses could not be detected
during the early biotrophic phase of U. maydis infection, indicating repressed SA
signaling (Seo et al. 2001; Doehlemann et al. 2008b). In summary, these studies
suggest that, once a biotrophic interface is established, U. maydis suppresses
defense responses and in particular HR-induced cell death.
Besides its ability to suppress defense mechanisms of the host plant a second
process that characterizes the biotrophic U. maydis infection is the reprogramming
of the plant physiology. So far detailed metabolic studies revealed a redirection of
carbohydrate and nitrogen fluxes towards the infected tissue in which massive
proliferation of fungal hyphae occurs (Horst et al. 2008; Doehlemann et al.
2008b; Horst et al. 2010).
Measurements of hexoses and hexose/sucrose ratios revealed that about
4–6 days post infection infected leaf areas convert from a carbon source to a sink
tissue (Billet and Burnett 1978; Doehlemann et al. 2008b; Horst et al. 2008). In line
Compatibility in Biotrophic Plant–Fungal Interactions: Ustilago maydis and Friends
221
with this, transitory starch production and sucrose accumulation during the light
period were dampened in tumor tissue.
Maize is a C4 plant. However, in the immature stage seedlings still exhibit C3 or
C3–C4 intermediate photosynthesis (Nelson and Dengler 1992; Langdale and
Kidner 1994). During early leaf development, uninfected juvenile maize seedlings
undergo a transition to C4 metabolism. In contrast, characteristic features of C3
photosynthesis are kept throughout U. maydis infections of maize seedlings,
demonstrating that the transition to C4 metabolism is prevented in infected
seedlings (Horst et al. 2008; Doehlemann et al. 2008b). U. maydis does not infect
differentiated source leaf tissue (Wenzler and Meins 1987). This suggests that
during infections of immature seedlings this fungus reprograms the developing
tissue in a way that the sink status is retained rather than transforming source leaves
back to sink leaves.
Stable isotope probing experiments demonstrated that U. maydis tumors furthermore constitute strong nitrogen sinks that rely on organic nitrogen import from
noninfected, systemic source leaves (Horst et al. 2010). In accordance with this,
Horst et al. (2010) provided evidence that the systemic source leaves display an
enhanced photosynthetic capacity as well as an increased amino acid export. These
amino acids are transported via the phloem to the tumor tissue where they are
metabolized to nitrogen-rich amino acids. These data suggest that the increased
import of nitrogen into tumor tissue might feed the protein biosynthesis of U.
maydis (Horst et al. 2010).
Recently, the identification of a novel sucrose transporter essential for pathogenicity improved our understanding of how U. maydis extracts nutrients from its host
(Wahl et al. 2010a). This specific transporter allows the direct utilization of sucrose
at the plant–fungal interface without extracellular hydrolysis and, thus, avoids the
production of extracellular monosaccharides known to elicit plant immune
responses (Wahl et al. 2010a). This finding also provides one example of the
means U. maydis employs to prevent the excessive elicitation of plant responses
during biotrophy.
5 Surface Sensing and Penetration
Initiation of different prepenetration stages on the plant surface is a prerequisite for
the successful establishment of biotrophy. These stages involve cell attachment, the
exploitation of suitable penetration sites, and the subsequent differentiation of
penetration structures. This requires a complex molecular dialogue between fungus
and plant (Dixon and Lamb 1990; Apoga et al. 2004).
On the plant surface, U. maydis undergoes a dimorphic switch from budding to
hyphal growth. Once compatible haploid sporidia recognize each other via a
pheromone–receptor system they form conjugation tubes that fuse at their tips.
This generates a dicaryotic filament – the infections form of U. maydis (Sleumer
1932; Bowman 1946; Rowell 1955; Snetselaar and Mims 1992; Snetselaar 1993;
222
K. Schipper and G. Doehlemann
Spellig et al. 1994). Attachment of U. maydis filaments to the hydrophobic plant
surface is likely mediated by repellent peptides. The rep1 gene encodes a secreted
precursor protein that is proteolytically cleaved into 11 peptides with high sequence
similarity during secretion (W€osten et al. 1996; Teertstra et al. 2006). These
peptides have functionally replaced hydrophobins in U. maydis and are likely to
function as amyloid-like structures on the hyphal surface (Teertstra et al. 2009).
About 16 h post infection fungal filaments growing on the plant surface differentiate appressoria that enable the fungus to enter the plant tissue and therefore,
initiate the switch to invasive growth (Snetselaar and Mims 1992; MendozaMendoza et al. 2009). The differentiation of filaments as well as appressoria in
U. maydis is dependent on host-derived physicochemical cues that are perceived
by the fungus (Mendoza-Mendoza et al. 2009).
The plant cuticle consists of two layers. The uppermost layer which is in direct
contact with the environment is a continuous layer of epicuticular waxes. The
second, inner layer is composed of intracuticular waxes associated with a polyester
matrix of cutin (Eigenbrode and Espelie 1995; Kolattukudy 2001). Cutin itself
resembles a network of interesterified hydroxyl and epoxy derivatives of C16 and
C18 fatty acids (Purdy and Kolattukudy 1975). Mendoza-Mendoza et al. (2009)
demonstrated that hydrophobicity induces the formation of dicaryotic U. maydis
filaments on artificial surfaces that resemble those growing on the surface of
infected plants. In the natural environment, this hydrophobic surface is most likely
provided by the plant cuticle.
An additional chemical cue, i.e., addition of hydroxy fatty acids, further
stimulates filament formation and moreover, induces appressorium differentiation
in the in vitro system. In addition, cutin monomers induce differentiation of
appressoria at similar rates as observed for hydroxy fatty acids. It has been
postulated that filamentously growing cells on the plant surface locally degrade
plant cutin by the secretion of lytic enzymes leading to the release of cutin
monomers. This may provide the stimulus that induces the differentiation of
appressoria (Mendoza-Mendoza et al. 2009).
U. maydis appressoria are nonmelanized and are therefore unlikely to function
through mechanical turgor force (Snetselaar and Mims 1993; Brachmann et al.
2003). This hypothesis is supported by the finding that during penetration, the
cytoplasm can be located in the hyphal part that is still on the leaf surface as well
as in the hyphal peg that already has penetrated (Schirawski et al. 2005).
Rather than employing mechanical force, U. maydis synthesizes a minimal set of
secreted lytic enzymes that enables the fungus to penetrate (Mueller et al. 2008;
Doehlemann et al. 2008a). Four secreted cutinases might degrade the cuticle that
covers the leaf surface (Mueller et al. 2008). Beneath the cuticle the maize cell wall
consists of glycan-crosslinked cellulose (Carpita et al. 2001; Abedon et al. 2006).
Penetration of this rigid barrier is likely mediated by the rather limited set of plantpolysaccharide degrading enzymes which partially degrade and soften the plant cell
wall to allow entry of the infectious filament (Mueller et al. 2008; Doehlemann
et al. 2008a; K€amper et al. 2006). At the same time, the poor equipment with cell
Compatibility in Biotrophic Plant–Fungal Interactions: Ustilago maydis and Friends
223
wall degrading enzymes might prevent the excessive generation of molecules that
lead to a stimulation of the plant defense machinery.
In contrast to the evident dome-shaped, melanized appressoria of pathogens like
M. grisea the corresponding nonmelanized structures in U. maydis are rather hard to
distinguish from other morphological structures. Characteristic but hardly recognizable features of filaments that form appressoria are slightly swollen tips which
directly emerge from hyphal bends. Nevertheless, the recent establishment of an
appressorial marker that consists of a gfp reporter gene fused to a promoter that is
specifically induced in hyphal tip cells differentiating appressoria provides a
sophisticated method to characterize differentiation and function of penetration
structures in U. maydis (Mendoza-Mendoza et al. 2009).
6 Effectors
Plants employ a basal PAMP-induced defense system that recognizes pathogen
attacks and subsequently triggers a defense program that includes an HR. Since an
HR finally results in the PCD of the attacked cell this comprises a highly efficient
way to block biotrophic pathogens. In the need to overcome this mechanism,
pathogens developed specialized secreted effectors that counteract plant defense
responses and thereby ensure compatibility (Jones and Dangl 2006).
A vast array of effectors has been identified in the past years of research. The
described proteins show a high structural and functional variety. However, these
proteins usually do not classify into known functional categories, i.e., enzyme
classes but rather represent completely novel proteins with no homologies to
proteins with described functions. Moreover, the secreted effectors often lack
known functional domains. Therefore, research on these unique proteins is particularly challenging. Especially, detailed functional studies of fungal effectors are still
scarce.
The question of effector function in biotrophic interactions is tightly intertwined
with the localization of these proteins. As classical effectors contain an N-terminal
signal peptide they are targeted to the secretory pathway of the pathogen and thus,
end up in the biotrophic interface after secretion (Fig. 1). Three main scenarios of
effector function were described until now: A variety of effector proteins inhibits
plant PR proteins (Tian et al. 2004; Rooney et al. 2005; Fig. 1). Such function could,
e.g., be assigned to the Avr2 effector of Cladosporium fulvum. This effector of the
causal agent of tomato leaf mold is able to inhibit cysteine proteases of the host
tomato (Rooney et al. 2005; van Esse et al. 2008). The hemibiotrophic oomycete P.
infestans also secretes effectors that guard the pathogen through the inhibition of
host proteases (Tian et al. 2004, 2007). U. maydis secretes a peroxidase that is likely
to detoxify H2O2 during the initial phase of plant infection (Molina and Kahmann
2007). Moreover, secreted effectors can mask the pathogen and thereby prevent
recognition by the host cell (Fig. 1). The chitin-binding virulence factor Avr4 of C.
fulvum covers the fungal cell wall during plant infection and thereby, impedes the
224
K. Schipper and G. Doehlemann
action of chitinases (Westerink et al. 2002; van den Burg et al. 2006; van Esse et al.
2007). Recently, the LysM-domain containing effector Ecp6 of C. fulvum has been
shown to prevent chitin-triggered immunity by sequestering chitin oligomers that
are released from the cell walls of invading hyphae (de Jonge et al. 2010). Another
reasonable function of pathogen effectors might be mediated by the binding to plant
receptors that, e.g., trigger defense responses (Fig. 1). Hints for such functions are
described interactions of secreted effectors with membrane-bound resistance
proteins. In many cases these interactions are indirect, following the guard hypothesis (Dangl and Jones 2001; van der Hoorn and Kamoun 2008) – in other cases
direct effector–receptor interactions have been demonstrated. An example is the
C. vulvum Avr4 protein, which directly binds to the cognate Cf-4 receptor in tomato
(van Esse et al. 2007).
In the last years it became more and more evident that secreted effectors of
(hemi)-biotrophic pathogens not only reside in the biotrophic interface. Intriguingly, some effectors can enter plant cells (Fig. 1) where they usually directly
interfere with defense-related processes (Ellis et al. 2009; Hogenhout et al. 2009;
Stergiopoulos and de Wit 2009). A multiplicity of translocated pathogen effectors
has been identified in phytopathogenic bacteria like X. campestris as well as in
oomycetes. While the bacterial pathogens use a type III or IV secretion system to
directly inject their effectors into the host cell a specific translocation motif (RXLR)
has been identified in the N-terminal region of transferred oomycete effector
molecules (Lahaye and Bonas 2001; Axtell et al. 2003; Gosh 2004; Kamoun
2007; Morgan and Kamoun 2007; Whisson et al. 2007). For phytopathogenic
fungi, translocation into the plant cell has been shown for the M. grisea effector
AVR-Pita, Uromyces fabae RTP1, and the flax rust effectors AvrM, AvrL657,
AvrP123, and AvrP4 (Jia et al. 2000; Kemen et al. 2005; Catanzariti et al. 2006;
Dodds et al. 2006; Rafiqi et al. 2010).
Plants potentially develop specific resistance (R) proteins that recognize pathogen effectors that interfere with defense responses. This in turn leads to effectortriggered immunity that is often associated with PCD induction (Jones and Dangl
2006). Remarkably in the U. maydis–maize interaction, such gene-for-gene
systems, i.e., effectors that are specifically recognized by cognate resistance
genes of the plant, have not been described so far. Possibly such an interaction
has not been developed in this pathosystem, probably because the pathogenic
pressure of U. maydis is not sufficient. An alternative explanation is that resistance
against U. maydis has not been observed in maize plants because a comprehensive
screen for resistance loci has not been performed yet.
In the recent past, the first effector proteins that actively interfere with PCD
induction have been described in plant pathogenic bacteria and oomycetes. One
example is the effector protein AvrPtoB of the plant pathogenic bacterium Pseudomonas syringae that is translocated into host cells (Rosebrock et al. 2007). The
oomycete Phytophtora infestans encodes AVR3a which has been shown to suppress cell death in Nicotinia benthamiana (Bos et al. 2006); the Phytophtora sojae
effector Avr1b facilitates suppression of BAX-induced cell death via action of
Compatibility in Biotrophic Plant–Fungal Interactions: Ustilago maydis and Friends
225
conserved motifs (Dou et al. 2008). For fungal effectors, a direct interference with
PCD induction could not be proven so far.
Sequencing of the U. maydis genome revealed a set of 386 genes encoding novel
proteins that are predicted to be secreted. Another 158 genes were predicted to code
for secreted enzymes (Mueller et al. 2008). Proteins of the first class are likely to
resemble secreted effectors that may have specific functions during biotrophic
development of U. maydis. Remarkably, many of the genes for novel secreted
proteins are arranged in clusters containing up to 24 genes (K€amper et al. 2006).
During biotrophic development, the majority of these clustered, putative effector
genes are transcriptionally up-regulated (K€amper et al. 2006). Twelve of these gene
clusters were deleted in the solopathogenic U. maydis strain SG200 and five of the
resulting mutants were significantly altered in virulence. While mutants of four
gene clusters were attenuated in virulence and displayed defects at different stages
of pathogenic development, one cluster mutant showed an increased virulence
(K€amper et al. 2006). In addition to genes encoding secreted proteins that are
arranged in clusters, U. maydis also comprises gene families where the individual
genes are dispersed upon the different chromosomes. Recently, the development of
a FLP-mediated marker recycling system enabled the analysis of such gene families
in U. maydis. The complete deletion of an 11 gene family (eff11) resulted in a
significant reduction of virulence (Khrunyk et al. 2010). This attenuation could be
attributed to three of the genes. Another family of effectors is encoded by the mig2
genes that were identified because of their strong up-regulation during plant
colonization (Basse et al. 2002). Five of the mig2 genes are clustered and a sixth
gene is located independently on the genome (Basse et al. 2002; Farfsing et al.
2005). All mig2 encoded proteins contain eight conserved cysteine residues, as it
also has been found in Avr proteins of C. fulvum (Stergiopoulos and de Wit 2009).
However, a mutant in which all six mig2 genes were deleted showed no reduction in
virulence when infected in maize seedlings. Given the high expression levels of
these genes, one might speculate on putative functions of these effectors. So far, the
mutants have only been tested in seedling infections of the maize variety Early
Golden Bantam. Therefore, one possibility is that the function of the Mig2 protein
family in virulence of U. maydis depends on the maize variety or the plant organ
being infected. Particularly, a tissue-specific function of these proteins would be of
interest, as recent findings suggest a similar situation for other U. maydis effectors
as well (see Sect. 8; Skibbe et al. 2010).
A single secreted effector that is not arranged in a gene cluster encoding other
secreted proteins has recently been shown to be essential for establishment of the
biotrophic interaction of U. maydis with maize plants. This protein, termed Pep1
(Protein essential during penetration 1), is dispensable for saprophytic growth of
U. maydis but deletion mutants are completely blocked in biotrophic development
(Doehlemann et al. 2009). Confocal microscopy showed the mutant being able to
penetrate the plant cell wall but subsequent invasion of the host cell stops immediately after invagination of the plant plasma membrane. At the same time, the mutant
induces strong plant defense responses including PCD of attacked epidermis cells.
Using fluorescently tagged versions as well as immunolocalization, the protein was
226
K. Schipper and G. Doehlemann
shown to be secreted from intracellular hyphae into the biotrophic interface,
particularly accumulating when hyphae invade a so far uncolonized cell. However,
the molecular function of Pep1 remains unclear. Since a Pep1 homolog of the barley
covered smut fungus Ustilago hordei was able to complement U. maydis deletion
mutants it is likely that it holds a general, conserved mechanism that is essential for
pathogenicity of smuts (Doehlemann et al. 2009). Currently, biochemical
approaches are applied to determine the molecular mechanism of Pep1. Given the
fundamental role of this effector for pathogenicity, its plant targets are supposed to
constitute major resistance factors of monocotyledonous plants to biotrophic fungi.
7 Regulation of Biotrophic Development
During biotrophic development, U. maydis undergoes a variety of stages that need
to be tightly regulated to assure compatibility. Characterization of the underlying
molecular mechanisms is essential to improve our understanding of how the fungus
is able to succeed in the intimate relationship with its host plant.
The initial step during pathogenic development of U. maydis is the mating of two
compatible sporidia on the plant surface (B€olker et al. 1992; Spellig et al. 1994).
Cell fusion leads to the formation of an active transcription factor termed bE/bW,
an heterodimeric protein that resembles a crucial cell cycle regulator. The active
heterodimer induces a G2 cell cycle arrest that is maintained during filamentous
growth on the leaf, during appressorium differentiation and during penetration of
the host tissue. Directly after penetration, as soon as a compatible biotrophic
interaction has been established, the G2 cell cycle arrest is released and filaments
start to proliferate intra- and intercellularly in the host tissue (Garcı́a-Muse et al.
2003; Scherer et al. 2006; Cánovas and Pérez-Martı́n 2009). bE/bW expression is a
prerequisite for the transition from saprophytic to biotrophic growth of U. maydis
since the b-heterodimer is required and sufficient to initiate pathogenic development and sexual reproduction (B€olker et al. 1995).
The essential role of the bE/bW transcription factor for pathogenic development
could be characterized using a strain harboring a mutant allele that encodes a
temperature-sensitive bE protein. This protein is instable at restrictive temperature.
Plant pathogenicity assays under these conditions revealed a complete block of
pathogenic development due to a failure in colonization of the host plant (Wahl
et al. 2010b). Detailed microscopic analyses demonstrated that in planta hyphae
develop severely enlarged, bulbous hyphal tip cells containing multiple nuclei,
indicating a defect in cell cycle control and cytokinesis (Wahl et al. 2010b).
Furthermore, transcriptional profiling under restrictive conditions conducted by
DNA microarrays revealed a down-regulation of genes encoding secreted effectors
during in planta growth. Intriguingly, the expression of 12 genes located in five U.
maydis gene clusters encoding secreted effectors depends on b-activation and 10 of
these belong to clusters that have been proven to execute important functions during
pathogenicity (Wahl et al. 2010b; K€amper et al. 2006; see Sect. 3). These results
Compatibility in Biotrophic Plant–Fungal Interactions: Ustilago maydis and Friends
227
further demonstrated the importance of the bE/bW heterodimer as the central
regulator of pathogenic development: The transcription factor is not only crucial
for the establishment of the dikaryon and for initiation of biotrophy but also for its
sustainment until sexual reproduction is completed (Wahl et al. 2010b).
To detect genes that are directly influenced by the bE/bW transcription factor
with respect to their regulation the b-genes were hooked up to inducible promoters.
Shifting of the U. maydis from restrictive to inductive growth conditions in axenic
culture leads to a synchronized synthesis of the b-heterodimier. This allows comprehensive view on b-regulated gene expression by DNA microarray analyses
(Heimel et al. 2010a). To this end, 347 b-regulated genes were identified, with a
subset of 212 genes up- and the remaining 135 genes down-regulated (Heimel et al.
2010a). In sum, transcripts encoding proteins that are involved in cell wall
remodeling, lipid metabolism, cell cycle control, mitosis, and DNA replication
were up-regulated. Moreover, a transcriptional induction could be detected for
genes that code for putative secreted effector proteins (Heimel et al. 2010a).
Interestingly, only 14 of the 345 b-dependently regulated genes that have been
identified by this transcriptional profiling monitoring b induction in axenic culture
were also differentially expressed after temperature restriction of b activity in
planta. This suggests that additional plant-specific signals and regulators modify
b-mediated transcription (Wahl et al. 2010b).
One example for a directly b-activated gene that plays an important role during
pathogenic development is Clp1. A Clp1 homolog has been shown to be essential
for clamp cell formation in Coprinus cinereus (Inada et al. 2001). U. maydis clp1
deletion strains are neither disturbed in appressorium formation nor in appressorium function. However, after penetration hyphal growth is arrested prior to the first
mitotic division (Scherer et al. 2006). This indicates that clp1 is required for
proliferation in the host tissue. Further investigations revealed that clp1 deletion
leads to a failure in the formation of clamp-like structures during invasive growth as
it has been described for the deletion of clp1 in C. cinereus. As a consequence, in
planta hyphae of clp1 deletion mutants display a disturbed nuclear distribution
(Scherer et al. 2006). While overexpression of clp1 in haploid U. maydis strains
does not lead to any obvious defects, overexpression of clp1 in strains with an
active bE/bW heterodimer strongly attenuates filamentation. Most likely, this
observation is due to the fact that Clp1 counteracts the function of the bE/bW
heterodimer. This is likely to occur via protein–protein interactions since the
expression of the b genes is not affected by clp1 expression. In line with this
hypothesis it could be shown that a set of b-induced genes is down-regulated in
clp1 overexpressor strains (Scherer et al. 2006). Presumably, a Clp1-mediated
transient inhibition of the bE/bW complex is needed to facilitate cell cycle progression during hyphal growth (Scherer et al. 2006).
Remarkably, most of the genes that were found to be transcriptionally controlled
by the active bE/bW heterodimer do not possess putative bE/bW-binding sites in
their promoter regions. Instead, it has been shown that a limited set of directly binduced (so-called class I b targets) transcription factors are responsible for the
regulation of the majority of b-regulated genes (90%; Heimel et al. 2010a). Thus,
228
K. Schipper and G. Doehlemann
the bE/bW protein does not directly influence the expression of all 347 target genes
but rather triggers a complex multilayered regulatory network.
One particular class I target gene involved in regulation of downstream genes, the
transcriptional regulator Rbf1, constitutes a C2H2 zinc finger protein. This transcription factor resembles the master regulator within the network of b-controlled genes
during pathogenic development (Heimel et al. 2010b). Strains deleted in the rbf1
allele display a similar phenotype as strains lacking an active bE/bW complex.
Complementary, induction of rbf1 leads to filament formation as well as a G2 cell
cycle arrest, as it is also observed after activation of the bE/bW heterodimer (Heimel
et al. 2010b).
The class I transcription factor Rbf1 controls another subset of regulators, the
class II transcription factors. Two of them, Biz1 and Hdp1, are important during
biotrophy (Heimel et al. 2010b; Spanu and K€amper 2010). Interestingly, Biz1
seems to regulate distinct stages of pathogenic development: The protein is
dispensable for mating and filamentation but biz1 deletion strains are strongly
disturbed in appressoria formation and in planta proliferation. Only few hyphae
are able to penetrate the plant epidermis and subsequent hyphal growth is stalled
directly (Flor-Parra et al. 2006). Although biz1 deletion strains display a very early
pathogenicity defect, the protein might be essential during the complete pathogenic
development since biz1 expression is up-regulated during all stages of biotrophy
(Flor-Parra et al. 2006).
Biz1 represses the expression of the cyclin clb1, resulting in a G2 arrest. Based
on this it was speculated that the Biz1-induced cell cycle arrest is important though
probably not sufficient for appressorium formation (Flor-Parra et al. 2006). The fact
that biz1 is highly expressed during the entire infection process might indicate that
U. maydis cells growing within the plant tissue continuously need to go through cell
cycle arrest stages.
U. maydis strains where genes for secreted effectors were deleted did not only
display phenotypes that are due to an early block of biotrophic development. Some
effector mutants showed reduced virulence and other rather subtle phenotypes that
can be traced back to defects during later stages of plant colonization (K€amper et al.
2006). On these grounds it has been speculated that gene expression during all
stages of pathogenicity is subject to a tight spatial and temporal regulation. Thus,
there is a need for regulatory transcription factors that specifically control in planta
gene expression. Recently, one such transcription factor, Mzr1, could be identified
in a combined approach of promoter analysis and expression profiling (Zheng et al.
2008). This zinc-finger transcription factor controls, e.g., the in planta expression of
three well-described genes that are specifically up-regulated during maize colonization. The three genes are part of the so-called mig2-gene cluster that consists of
five related genes (mig2-1 to mig2-5) coding for secreted proteins with yet unknown
functions (Basse et al. 2002). Moreover, it has been demonstrated that the promoter
region of one cluster gene, mig2-5, contains a cis-active element that mediates a
strong transcriptional activation by Mzr1 during biotrophic development (Farfsing
et al. 2005; Zheng et al. 2008). Although DNA microarray analyses revealed around
50 Mzr1-induced U. maydis transcripts with an enrichment of transcripts encoding
Compatibility in Biotrophic Plant–Fungal Interactions: Ustilago maydis and Friends
229
secreted proteins, deletion of mzr1 only leads to reduced pathogenicity (Zheng et al.
2008). This result demonstrates again that regulation of pathogenic development
cannot be traced back to a single transcription factor but that biotrophy is controlled
by a complex network of transcription factors. Moreover, these transcriptional
regulators seem to be tightly cross-linked as illustrated by the example of the
interdependence between Mzr1 and Biz1: Although Mzr1 itself is independent
from Biz1, activity of Biz1 was found to be necessary for expression of the
mig2–6 gene, which in turn is directly depending on Mzr1.
Another example of biotrophy-specific transcription factors in U. maydis is the
forkhead protein Fox1. This protein is the first example of a b-independent transcription factor, indicating that biotrophic development is not only regulated by the
b-network but also includes other important factors (Zahiri et al. 2010). Plants
infected with fox1 deletion mutants display enhanced defense reactions and therefore show drastically reduced virulence. This proofs the impact of b-independent
regulators in U. maydis (Zahiri et al. 2010). Transcriptional profiling revealed about
130 U. maydis genes lacking transcriptional induction in the fox1 deletion strain and
these were again enriched for genes that code for putative secreted effectors
(33 genes). However, single-gene deletion mutants for some of the most downregulated U. maydis genes did not result in reduced virulence like observed for fox1
deletion mutants. Thus, it may be speculated that only the simultaneous deletion of
several or a distinct combination of Fox1-induced genes may mirror the fox1
phenotype (Zahiri et al. 2010).
8 Stage and Organ Specificity of U. maydis Effectors
U. maydis virulence underlies complex regulatory networks during disease progression. However, it has recently been demonstrated for the first time that for establishment of biotrophy spatial regulatory fine tuning of infection seems to be of equal
importance (Skibbe et al. 2010).
From the time when penetration occurs, U. maydis actively suppresses basal
PAMP-triggered plant immune responses (Doehlemann et al. 2008b; see Sect. 4).
However, biotrophy also includes active long-term manipulation of the host. Thus,
it is not sufficient for the fungus to suppress only early plant defense responses.
Secreted effectors which are known to be crucial for biotrophy are likely deployed
for both processes. In accordance with this, there is experimental evidence that
Pep1, an effector that is a major player in suppression of plant defense responses, is
not only essential for the initial penetration of the plant surface but also for
subsequent cell-to-cell passages in the maize tissue (Doehlemann et al. 2009).
Along with the analysis of gene clusters for secreted effectors in U. maydis also
cluster deletion mutants with “mild” phenotypes due to defects during later stages of
infection were identified. Observed phenotypes of cluster deletion mutants that did
not completely loose virulence range from slight to strong reductions in tumor size
and number to hypervirulence (K€amper et al. 2006). Though mutants like, e.g., the
230
K. Schipper and G. Doehlemann
cluster 19a deletion mutant are able to invade the plant tissue, tumor induction can be
observed only very rarely (K€amper et al. 2006). These diverse phenotypes implicate
a stage-specific activity of some secreted effectors. It seems obvious that U. maydis
needs to employ specific sets of effectors during the different phases of infection
since the requirements on the host will change with the course of infection that
involves penetration, intracellular growth, and massive proliferation in intercellular
spaces. The probably most elementary host cell reprogramming events need to occur
during tumor formation – a process that is rarely understood to date. Unraveling the
molecular mechanisms underlying these manipulations of the host by U. maydis
effectors will be one particularly exciting aspect of future studies.
A unique feature of U. maydis in comparison to other, closely related smut fungi
is its ability to induce tumor and subsequent spore formation in basically all aerial
parts of its host plant. In contrast, symptom formation in infections with other smut
fungi like, for instance, the head smut Sporisorium reilianum is restricted to the
inflorescences of the respective host plants. With its ability to colonize a variety of
maize tissues encountering cells with dissimilar grades of differentiation, U. maydis
must be able to tailor its effector weaponry to these specific conditions (Skibbe et al.
2010). During ontogeny as well as in the mature form, each maize tissue produces a
particular combination of proteins. Thus, it seems reasonable that multiple maize
quantitative trait loci were identified which modulate the extent of tumor development in specific organs like the tassel, stalk, or ear tissue during U. maydis infection
(Baumgarten et al. 2007).
With a simultaneous transcriptional profiling of U. maydis and Z. mays genes
during the biotrophic stage, huge differences in the transcriptome of different
infected host organs (seedling leaf, adult leaf, and tassel) were identified. The
results indicated that the transition of leaf tissue to the convenience of the fungus
requires significantly more changes in gene expression than in the tassel. Interestingly, not only the total number of regulated transcripts but also the respective
functions of plant genes differentially expressed during U. maydis infection
diverged between individual host organs (Skibbe et al. 2010). The results indicate
that the special attribute of U. maydis to form tumors not only in the flowers but also
on leaves must have been developed secondarily. In contrast, other smut fungi did
not gain this competency. Moreover, the study proofed that mutants of the gene
clusters encoding secreted effectors also display organ-specific phenotypes. The
most intriguing example for this specificity is the cluster 19a deletion mutant: The
corresponding strains induce tumor formation only very rarely in seedling leaves
where the cells showed huge transcriptional changes during tumor induction.
Contrarily, cluster 19a mutants behave almost like wild-type strains when tassels
are infected, suggesting that the corresponding effectors are not crucial for symptom development in the male inflorescence (Skibbe et al. 2010). It will be a future
challenge to elucidate which functions tissue-specific virulence factors exhibit
during transformation of normal maize cell fates according to the specific
requirements of the fungus in the respective organ of the plant.
According to the state of our knowledge, we conclude that the biotrophic
interaction between U. maydis and its host Z. mays most likely consists of two
Compatibility in Biotrophic Plant–Fungal Interactions: Ustilago maydis and Friends
231
major phases: Initially, compatibility is established by general pathogenicity factors
that suppress the PAMP-elicited basal plant defense responses during penetration
(Doehlemann et al. 2008b). In a second phase of infection, fungal proliferation in
the respective infected plant tissue is assured by specific subsets of effectors that are
expressed in a strictly regulated spatial-temporal manner. Thus, the fungus is able to
efficiently reprogram plant physiology and development of a specific organ tissue.
It is tempting to speculate whether such a strategy is also deployed by other
pathogenic organisms that infect plants as well as animal hosts.
9 Perspectives
The enormous progress made by the research community within the last years
provided a substantial body of knowledge on the pathogenic development of fungal
biotrophs and the regulatory mechanisms controlling these processes. A key role in
establishment and maintenance of biotrophy is held by secreted effector proteins
that directly interfere with plant targets and thereby control host defenses and
metabolic reprogramming. In the U. maydis–maize pathosystem, the combination
of genome sequencing, transcriptome profiling, and powerful reverse genetics
identified a number of effectors with essential functions at distinct steps of pathogenic development. However, the molecular mechanisms of those effectors, their
subcellular localization within the plant tissue as well as the cellular processes of
the plant they target have not been clarified so far. For this reason, a major task for
the near future will be to functionally analyze the U. maydis effectors that have
important functions during the biotrophic interaction. This will require a smart
combination of molecular, biochemical, and microscopic technologies together
with bioinformatics resources. Comparative genomics of related smut fungi will
be an important tool to identify conserved motifs in effectors that might give hints
on their function as well as their localization. Effectome comparison in different
smuts will also allow discriminating between host-specific proteins and factors
being generally required for biotrophic interaction.
To study effector function in U. maydis, a major challenge will be to develop
transient protein expression systems in maize that are compatible with the fungal
infection. In other systems such as the B. graminis–barley interaction, both for
transient overexpression and gene silencing, particle bombardment is an excellent
tool to study the role of host genes on pathogen interaction (Eichmann et al. 2004).
These assays also provide the possibility of a functional characterization of fungal
effectors and their interactions with the respective plant targets. Since smut fungi
only infect meristematic tissue, this kind of assay cannot be used to directly
determine effects on fungal pathogenicity. The systemic VIGS system that recently
has been described is a first step to allow a rapid functional characterization of
putative effector-interacting maize proteins (van der Linde et al. 2010).
Candidate genes for effectors as well as for plant factors involved in the
establishment and maintenance of compatibility often are selected on the basis of
232
K. Schipper and G. Doehlemann
transcript profiling approaches. However, all microarray data on the U.
maydis–maize interaction has been gained from infected sections of whole plant
organs, i.e., leaves and tassel. Particularly in the early stages of interaction only a
low percentage of plant cells are actually in contact with fungal hyphae. In turn, the
fungal material represents only a small fraction (<5%; Horst et al. 2010) in samples
of infected leaves. This situation “dilutes” cell-specific effects and particularly
subtle alterations in gene expression might be overlooked so far. Therefore, dissection of individual cells from different tissues will provide a much more detailed
picture on the molecular processes associated with the intimate biotrophic interaction of U. maydis and its host.
Technical advances in confocal microscopy, -omics technologies, and the
sequencing of reference genomes allowed completely new insights into biotrophic
plant–pathogen interactions in the recent past. Now, high-throughput screening of
effectors, comparative genome analysis, and in-depth transcriptome profiling based
on second- and third-generation sequencing technologies will open new windows to
unravel the hidden secrets of the friendliest of all antagonistic microbes – the
biotrophs.
Acknowledgements We are grateful to Regine Kahmann and the Max Planck Institute for
Terrestrial Microbiology for continuous support. Our research is funded by grants of the Deutsche
Forschungsgemeinschaft (DFG) via Research Group FOR666 and Priority Program SPP1212, the
Loewe Center SYNMIKRO, the EMBO STF program and the Deutsche Bundesstiftung Umwelt
(DBU).
References
Abedon BG, Hatfield RD, Tracy WF (2006) Cell wall composition in juvenile and adult leaves of
maize (Z. mays). J Agric Food Chem 54:3896–3900
Apoga D, Barnard J, Craighead HG, Hoch HC (2004) Quantification of substratum contact
required for initiation of Collectotrichum graminicola appressoria. Fungal Genet Biol 41:1–12
Axtell MJ, Chisholm ST, Dahlbeck D, Staskawicz BJ (2003) Genetic and molecular evidence that
the Pseudomonas syringae type III effector protein AvrRpt2 is a cysteine protease. Mol
Microbiol 49:1537–1546
Bailey JA, O’Connell JA, Pring RJ, Nash C (1992) Infection strategies of Colletotrichum species.
In: Bailey JA, Jeger MJ (eds) Colletotrichum: biology, pathology and control. CABI,
Wallingford, UK, pp 88–120
Banuett F (1995) Genetics of Ustilago maydis, a fungal pathogen that induces tumors in maize.
Annu Rev Genet 29:179–208
Banuett F, Herskowitz I (1996) Discrete developmental stages during teliospore formation in the
corn smut fungus, Ustilago maydis. Development 122:2965–2976
Basse CW (2005) Dissecting defense-related and developmental transcriptional responses of
maize during Ustilago maydis infection and subsequent tumor formation. Plant Physiol
138:1774–1784
Basse CW, Steinberg G (2004) Ustilago maydis, model system for analysis of the molecular basis
of fungal pathogenicity. Mol Plant Pathol 5:83–92
Basse CW, Kolb S, Kahmann R (2002) A maize-specifically expressed gene cluster in Ustilago
maydis. Mol Microbiol 43:75–93
Compatibility in Biotrophic Plant–Fungal Interactions: Ustilago maydis and Friends
233
Bauer R, Oberwinkler F, Vánky K (1997) Ultrastructural markers and systematics in smut fungi
and allied taxa. Can J Bot 75:1273–1314
Baumgarten AM, Suresh J, May G, Phillips RL (2007) Mapping QTLs contributing to Ustilago
maydis resistance in specific plant tissues of maize. Theor Appl Genet 114:1229–1238
Billet EE, Burnett JH (1978) The host-parasite physiology of the maize smut fungus Ustilago
maydis. 11. Translocation of 14C-labelled assimilates in smutted maize plants. Physiol Plant
Pathol 12:102–112
B€olker M, Urban M, Kahmann R (1992) The a mating type locus of U. maydis specifies cell
signaling components. Cell 68:441–450
B€olker M, Genin S, Lehmler C, Kahmann R (1995) Genetic regulation of mating and dimorphism
in Ustilago maydis. Can J Bot 73:329–342
Bos JI, Kanneganti TD, Young C, Cakir C, Huitema E, Win J, Armstrong MR, Birch PR, Kamoun S
(2006) The C-terminal half of Phytophthora infestans RXLR effector AVR3a is sufficient to
trigger R3a-mediated hypersensitivity and suppress INF1-induced cell death in Nicotiana
benthamiana. Plant J 48:165–176
Bowman DH (1946) Sporidial fusion in Ustilago maydis. J Agric Res 72:233–243
Brachmann A, Weinzierl G, K€amper J, Kahmann R (2001) Identification of genes in the bW/bE
regulatory cascade in Ustilago maydis. Mol Microbiol 42:1047–1063
Brachmann A, Schirawski J, M€
uller P, Kahmann R (2003) An unusual MAP kinase is required for
efficient penetration of the plant surface by Ustilago maydis. EMBO J 22:2199–2210
Brader G, Tas E, Palva ET (2001) Jasmonate-dependent induction of indole glucosinolates in
Arabidopsis by culture filtrates of the nonspecific pathogen Erwinia carotovora. Plant Physiol
126:849–860
Brefort T, Doehlemann G, Mendoza-Mendoza A, Reissmann S, Djamei A, Kahmann R (2009)
Ustilago maydis as a pathogen. Annu Rev Phytopathol 47:423–445
Cánovas D, Pérez-Martı́n J (2009) Sphingolipid biosynthesis is required for polar growth in the
dimorphic phytopathogen Ustilago maydis. Fungal Genet Biol 46:190–200
Carpita NC, Defernez M, Findlay K, Wells B, Shoue DA, Catchpole G, Wilson RH, McCann MC
(2001) Cell wall architecture of the elongating maize coleoptile. Plant Physiol 127:551–565
Catanzariti AM, Dodds PN, Lawrence GJ, Ayliffe MA, Ellis JG (2006) Haustorially expressed
secreted proteins from flax rust are highly enriched for avirulence elicitors. Plant Cell
18:243–256
Chinchilla D, Zipfel C, Robatzek S, Kemmerling B, N€
urnberger T, Jones JD, Felix G, Boller T
(2007) A flagellin-induced complex of the receptor FLS2 and BAK1 initiates plant defence.
Nature 448:497–500
Christensen JJ (1963) Corn smut induced by Ustilago maydis. Am Phytopathol Soc Monogr
2:1–41
Dangl JL, Jones JD (2001) Plant pathogens and integrated defence responses to infection. Nature
411:826–833
de Jonge R, van Esse HP, Kombrink A, Shinya T, Desaki Y, Bours R, van der Krol S, Shibuya N,
Joosten MH, Thomma BP (2010) Conserved fungal LysM effector Ecp6 prevents chitintriggered immunity in plants. Science 329:953–1055
de Wit PJ (2007) How plants recognize pathogens and defend themselves. Cell Mol Life Sci
64:2726–2732
di Stasio M, Brefort T, Mendoza-Mendoza A, M€
unch K, Kahmann R (2009) The dual specificity
phosphatase Rok1 negatively regulates mating and pathogenicity in Ustilago maydis. Mol
Microbiol 73:73–88
Dixon RA, Lamb C (1990) Molecular communication in interactions between plants and microbial
pathogens. Annu Rev Plant Mol Biol 41:339–367
Dodds PN, Lawrence GJ, Catanzariti AM, Teh T, Wang CI, Ayliffe MA, Kobe B, Ellis JG (2006)
Direct protein interaction underlies gene-for-gene specificity and coevolution of the flax
resistance genes and flax rust avirulence genes. Proc Natl Acad Sci USA 103:8888–8893
234
K. Schipper and G. Doehlemann
Doehlemann G, Wahl R, Vranes M, de Vries RP, K€amper J, Kahmann R (2008a) Establishment of
compatibility in the Ustilago maydis/maize pathosystem. J Plant Physiol 165:29–40
Doehlemann G, Wahl R, Horst RJ, Voll LM, Usadel B, Poree F, Stitt M, Pons-K€
uhnemann J,
Sonnewald U, Kahmann R, K€amper J (2008b) Reprogramming a maize plant: transcriptional
and metabolic changes induced by the fungal biotroph Ustilago maydis. Plant J 56:181–195
Doehlemann G, van der Linde K, Assmann D, Schwammbach D, Hof A, Mohanty A, Jackson D,
Kahmann R (2009) Pep1, a secreted effector protein of Ustilago maydis, is required for
successful invasion of plant cells. PLoS Pathog 5:e1000290
Dou D, Kale SD, Wang X, Chen Y, Wang Q, Wang X, Jiang RH, Arredondo FD, Anderson RG,
Thakur PB, McDowell JM, Wang Y, Tyler BM (2008) Conserved C-terminal motifs required
for avirulence and suppression of cell death by Phytophthora sojae effector Avr1b. Plant Cell
20:1118–1133
Eichmann R, Schultheiss H, Kogel KH, H€
uckelhoven R (2004) The barley apoptosis suppressor
homologue BAX inhibitor-1 compromises nonhost penetration resistance of barley to the
inappropriate pathogen Blumeria graminis f. sp. tritici. Mol Plant Microbe Interact 17:484–490
Eigenbrode SD, Espelie KE (1995) Effects of plant epicuticular lipids on insect herbivores. Annu
Rev Entomol 40:171–194
Ellis JG, Rafiqi M, Gan P, Chakrabarti A, Dodds P (2009) Recent progress in discovery and
functional analysis of effector proteins of fungal and oomycete plant pathogens. Curr Opin
Plant Biol 12:1–7
Farfsing JW, Auffarth K, Basse CW (2005) Identification of cis-active elements in Ustilago
maydis mig2 promoters conferring high-level activity during pathogenic growth in maize.
Mol Plant Microbe Interact 18:75–87
Farrar JF (1984) In: Wood RKS, Jellis GJ (eds) In plant, disease: infection damage and loss.
Blackwell, Oxford
Flor-Parra I, Vranes M, K€amper J, Pérez-Martı́n J (2006) Biz1, a zinc finger protein required for
plant invasion by Ustilago maydis, regulates the levels of a mitotic cyclin. Plant Cell
18:2369–2387
Francis SA, Dewey FM, Gurr SJ (1996) The role of cutinase in germling development and
infection by Erysiphe graminis f. sp. hordei. Physiol Mol Plant Pathol 49:201–211
Garcı́a-Muse T, Steinberg G, Pérez-Martı́n J (2003) Pheromone-induced G2 arrest in the phytopathogenic fungus Ustilago maydis. Eukaryot Cell 2:494–500
Glazebrook J (2005) Contrasting mechanisms of defense against biotrophic and necrotrophic
pathogens. Annu Rev Phytopathol 43:205–227
Gosh P (2004) Process of protein transport by the type III secretion system. Microbiol Mol Biol
Rev 68:771–795
Greenberg JT, Yao N (2004) The role and regulation of programmed cell death in plant–pathogen
interactions. Cell Microbiol 6:201–211
Haldar K, Kamoun S, Hiller BS, Ooij C (2006) Common infection strategies of pathogenic
eukaryotes. Nat Rev Microbiol 4:922–931
Hammond-Kosack KE, Parker JE (2003) Deciphering plant-pathogen communication; fresh
perspectives for molecular resistance breeding. Curr Opin Biotechnol 14:177–183
Heimel K, Scherer M, Schuler D, K€amper J (2010a) The Ustilago maydis Clp1 protein orchestrates
pheromone and b-dependent signaling pathways to coordinate the cell cycle and pathogenic
development. Plant Cell 22:2908–2922
Heimel K, Scherer M, Vranes M, Wahl R, Pothiratana C, Schuler D, Vincon V, Finkernagel F,
Flor-Parra I, K€amper J (2010b) The transcription factor Rbf1 is the master regulator for
b-mating type controlled pathogenic development in Ustilago maydis. PLoS Pathog 6:
e1001035
Hogenhout SA, Van der Hoorn RA, Terauchi R, Kamoun S (2009) Emerging concepts in effector
biology of plant-associated organisms. Mol Plant Microbe Interact 22:115–122
Horst RJ, Engelsdorf T, Sonnewald U, Voll LM (2008) Infection of maize leaves with Ustilago
maydis prevents establishment of C4 photosynthesis. J Plant Physiol 165:19–28
Compatibility in Biotrophic Plant–Fungal Interactions: Ustilago maydis and Friends
235
Horst RJ, Doehlemann G, Wahl R, Hofmann J, Schmiedl A, Kahmann R, K€amper J, Sonnewald U,
Voll LM (2010) Ustilago maydis infection strongly alters organic nitrogen allocation in maize
and stimulates productivity of systemic source leaves. Plant Physiol 152:293–308
Howard RJ, Ferrari MA, Roach DH, Money NP (1991) Penetration of hard substrates by a fungus
employing enormous turgor pressures. Proc Natl Acad Sci USA 88(24):11281–11284
Inada K, Morimoto Y, Arima T, Murata Y, Kamada T (2001) The clp1 gene of the mushroom
Coprinus cinereus is essential for A-regulated sexual development. Genetics 157:133–140
Jia Y, McAdams SA, Bryan GT, Hershey HP, Valent B (2000) Direct interaction of resistance
gene and avirulence gene products confers rice blast resistance. EMBO J 19:4004–4014
Jones JD, Dangl JL (2006) The plant immune system. Nature 444:323–329
Kahmann R, Steinberg G, Basse C, Feldbr€
ugge M, K€amper J (2000) Ustilago maydis, the causative
agent of corn smut disease. In: Kronstad JW (ed) Fungal pathology. Kluwer, Dodrecht, The
Netherlands, pp 347–371
Kamoun S (2007) Groovy times: filamentous pathogen effectors revealed. Curr Opin Plant Biol
10:358–365
K€amper J (2004) A PCR-based system for highly efficient generation of gene replacement mutants
in Ustilago maydis. Mol Genet Genomics 271:103–110
K€amper J, Kahmann R, B€
olker M, Ma LJ, Brefort T, Saville BJ, Banuett F, Kronstad JW, Gold SE,
M€uller O, Perlin MH, W€
osten HA, de Vries R, Ruiz-Herrera J, Reynaga-Peña CG, Snetselaar K,
McCann M, Pérez-Martı́n J, Feldbr€
ugge M, Basse CW, Steinberg G, Ibeas JI, Holloman W,
Guzman P, Farman M, Stajich JE, Sentandreu R, González-Prieto JM, Kennell JC, Molina L,
Schirawski J, Mendoza-Mendoza A, Greilinger D, M€
unch K, R€
ossel N, Scherer M, Vranes M,
Ladendorf O, Vincon V, Fuchs U, Sandrock B, Meng S, Ho EC, Cahill MJ, Boyce KJ, Klose J,
Klosterman SJ, Deelstra HJ, Ortiz-Castellanos L, Li W, Sanchez-Alonso P, Schreier PH,
H€auser-Hahn I, Vaupel M, Koopmann E, Friedrich G, Voss H, Schl€
uter T, Margolis J, Platt
D, Swimmer C, Gnirke A, Chen F, Vysotskaia V, Mannhaupt G, G€
uldener U, M€
unsterk€
otter M,
Haase D, Oesterheld M, Mewes HW, Mauceli EW, DeCaprio D, Wade CM, Butler J, Young S,
Jaffe DB, Calvo S, Nusbaum C, Galagan J, Birren BW (2006) Insights from the genome of the
biotrophic fungal plant pathogen Ustilago maydis. Nature 444:97–101
Kemen E, Kemen AC, Rafiqi M, Hempel U, Mendgen K, Hahn M, Voegele RT (2005) Identification of a protein from rust fungi transferred from haustoria into infected plant cells. Mol Plant
Microbe Interact 18:1130–1139
Kemmerling B, Schwedt A, Rodriguez P, Mazzotta S, Frank M, Qamar SA, Mengiste T,
Betsuyaku S, Parker JE, M€
ussig C, Thomma BP, Albrecht C, de Vries SC, Hirt H, N€
urnberger
T (2007) The BRI1-associated kinase 1, BAK1, has a brassinolide-independent role in plant
cell-death control. Curr Biol 17:1116–1122
Khang CH, Berruyer R, Giraldo MC, Kankanala P, Park SY, Czymmek K, Kang S, Valent B
(2010) Translocation of Magnaporthe oryzae effectors into rice cells and their subsequent cellto-cell movement. Plant Cell 22:1388–1403
Khrunyk Y, M€unch K, Schipper K, Lupas AN, Kahmann R (2010) The use of FLP-mediated
recombination for the functional analysis of an effector gene family in the biotrophic smut
fungus Ustilago maydis. New Phytol 187:957–968
uchel A (eds) Polyesters I: biological
Kolattukudy PE (2001) Cutin from plants. In: Doi Y, Steinb€
systems and biotechnological production, vol 3. Wiley, Weinheim, pp 1–35
Lahaye T, Bonas U (2001) Molecular secrets of bacterial type III effector proteins. Trends Plant
Sci 6:479–485
Langdale JA, Kidner CA (1994) Bundle-sheath defective, a mutation that disrupts cellulardifferentiation in maize leaves. Development 120:673–681
Latunde-Dada AO (2001) Colletotrichum: tales of forcible entry, stealth, transient confinement
and breakout. Mol Plant Pathol 2:187–198
Lewis DH (1973) Concepts in fungal nutrition and the origin of biotrophy. Biol Rev 48:261–278
Mendgen K, Deising H (1993) Infection structures of fungal plant pathogens – a cytological and
physiological evaluation. New Phytol 124:193–213
236
K. Schipper and G. Doehlemann
Mendgen K, Hahn M (2002) Plant infection and the establishment of fungal biotrophy. Trends
Plant Sci 7:352–356
Mendoza-Mendoza A, Berndt P, Djamei A, Weise C, Linne U, Marahiel M, Vranes M, K€amper J,
Kahmann R (2009) Physical-chemical plant-derived signals induce differentiation in Ustilago
maydis. Mol Microbiol 71:895–911
Molina L, Kahmann R (2007) An Ustilago maydis gene involved in H2O2 detoxification is
required for virulence. Plant Cell 19:2293–2309
Money NP, Howard RJ (1996) Confirmation of a link between fungal pigmentation, turgor
pressure and pathogenicity using a new method of turgor measurement. Fungal Genet Biol
20:217–227
Morgan W, Kamoun S (2007) RXLR effectors of plant pathogenic oomycetes. Curr Opin
Microbiol 10:332–338
Mueller O, Kahmann R, Aguilar G, Trejo-Aguilar B, Wu A, de Vries RP (2008) The secretome of
the maize pathogen Ustilago maydis. Fungal Genet Biol 45(Suppl 1):S63–S70
Nelson T, Dengler NG (1992) Photosynthetic tissue differentiation in C4 plants. Int J Plant Sci 153:
S93–S105
O’Connell RJ, Panstruga R (2006) Tête à tête inside a plant cell: establishing compatibility
between plants and biotrophic fungi and oomycetes. New Phytol 171:699–718
Oliver RP, Ipcho SVS (2004) Arabidopsis pathology breathes new life into the necrotrophs-vsbiotrophs classification of fungal pathogens. Mol Plant Pathol 5:347–352
Perfect ES, Green JR (2001) Infection structures of biotrophic and hemibiotrophic fungal plant
pathogens. Mol Plant Pathol 2:101–108
Pryce-Jones E, Carver T, Gurr SJ (1999) The roles of cellulase enzymes and mechanical force in
host penetration by Erysiphe graminis f. sp. hordei. Physiol Mol Plant Pathol 55:175–182
Purdy RE, Kolattukudy PE (1975) Hydrolysis of plant cuticle by plant pathogens. Purification,
amino acid composition, and molecular weight of two isozymes of cutinase and a non-specific
esterase from Fusarium solani f. pisi. Biochemistry 14:2824–2831
Rafiqi M, Gan PH, Ravensdale M, Lawrence GJ, Ellis JG, Jones DA, Hardham AR, Dodds PN
(2010) Internalization of flax rust avirulence proteins into flax and tobacco cells can occur in
the absence of the pathogen. Plant Cell 22:2017–2032
Rooney HC, Van’t Klooster JW, van der Hoorn RA, Joosten MH, Jones JD, de Wit PJ (2005)
Cladosporium Avr2 inhibits tomato Rcr3 protease required for Cf-2-dependent disease resistance. Science 308:1783–1786
Rosebrock TR, Zeng L, Brady JJ, Abramovitch RB, Xiao F, Martin GB (2007) A bacterial E3
ubiquitin ligase targets a host protein kinase to disrupt plant immunity. Nature 448:370–374
Rowell JB (1955) Functional role of compatibility factors and an in vitro test for sexual compatibility with haploid lines of Ustilago zeae. Phytopathology 45:370–374
Scherer M, Heimel K, Starke V, K€amper J (2006) The Clp1 protein is required for clamp formation
and pathogenic development of Ustilago maydis. Plant Cell 18:2388–2401
Schirawski J, B€ohnert HU, Steinberg G, Snetselaar K, Adamikova L, Kahmann R (2005) ER
glucosidase II is required for pathogenicity of Ustilago maydis. Plant Cell 17:1–12
Schulze-Lefert P, Panstruga R (2003) Establishment of biotrophy by parasitic fungi and
reprogramming of host cells for disease resistance. Annu Rev Phytopathol 41:641–667
Seo HS, Song JT, Cheong JJ, Lee YH, Lee YW, Hwang I, Lee JS, Choi YD (2001) Jasmonic acid
carboxyl methyltransferase: a key enzyme for jasmonate-regulated plant responses. Proc Natl
Acad Sci USA 98:4788–4793
Shrawat AK, L€orz H (2006) Agrobacterium-mediated transformation of cereals: a promising
approach crossing barriers. Plant Biotechnol J 4:575–603
Skibbe DS, Doehlemann G, Fernandes J, Walbot V (2010) Maize tumors caused by Ustilago
maydis require organ-specific genes in host and pathogen. Science 328:89–92
€
Sleumer HO (1932) Uber
Sexualit€at und Zytologie von Ustilago zeae. (Beckm.). Unger Z Botan
25:209–263
Snetselaar KM (1993) Microscopic observation of Ustilago maydis mating interactions. Exp
Mycol 17:345–355
Compatibility in Biotrophic Plant–Fungal Interactions: Ustilago maydis and Friends
237
Snetselaar KM, Mims CW (1992) Sporidial fusion and infection of maize seedlings by the smut
fungus Ustilago maydis. Mycologia 84:193–203
Snetselaar KM, Mims CW (1993) Infection of maize by Ustilago maydis: light and electron
microscopy. Phytopathology 83:843–850
Snetselaar KM, Mims CW (1994) Light and electron microscopy of Ustilago maydis hyphae in
maize. Mycol Res 98:347–355
Snetselaar KM, Boelker M, Kahmann R (1996) Ustilago maydis mating hyphae orient their growth
toward pheromone sources. Fungal Genet Biol 20:299–312
Spanu P, K€amper J (2010) Genomics of biotrophy in fungi and oomycetes-emerging patterns. Curr
Opin Plant Biol 13:409–414
Spellig T, B€olker M, Lottspeich F, Frank RW, Kahmann R (1994) Pheromones trigger filamentous
growth in Ustilago maydis. EMBO J 13:1620–1627
Spellig T, Bottin A, Kahmann R (1996) Green fluorescent protein (GFP) as a new vital Marker in
the phytopathogenic fungus Ustilago maydis. Mol Gen Genet 252:503–509
Staples RC (2000) Research on the rust fungi during the twentieth century. Annu Rev Phytopathol
38:49–69
Staples RC (2003) A novel gene for rust resistance. Trends Plant Sci 8:149–151
Stergiopoulos I, de Wit PJ (2009) Fungal effector proteins. Annu Rev Phytopathol 47:233–263
Tanaka A, Christensen MJ, Takemoto D, Park P, Scott B (2006) Reactive oxygen species play a
role in regulating a fungus-perennial ryegrass mutualistic interaction. Plant Cell 18:1052–1066
Tanaka A, Takemoto D, Hyon GS, Park P, Scott B (2008) NoxA activation by the small GTPase
RacA is required to maintain a mutualistic symbiotic association between Epichloe¨ festucae
and perennial ryegrass. Mol Microbiol 68:1165–1178
Teertstra WR, Deelstra HJ, Vranes M, Bohlmann R, Kahmann R, K€amper J, W€
osten HA (2006)
Repellents have functionally replaced hydrophobins in mediating attachment to a hydrophobic
surface and in formation of hydrophobic aerial hyphae in Ustilago maydis. Microbiology
152:3607–3612
Teertstra WR, van der Velden GJ, de Jong JF, Kruijtzer JA, Liskamp RM, Kroon-Batenburg LM,
M€uller WH, Gebbink MF, W€
osten HA (2009) The filament-specific Rep1-1 repellent of the
phytopathogen Ustilago maydis forms functional surface-active amyloid-like fibrils. J Biol
Chem 284:9153–9159
Tian M, Huitema E, Da Cunha L, Torto-Alalibo T, Kamoun S (2004) A Kazal-like extracellular
serine protease inhibitor from Phytophthora infestans targets the tomato pathogenesis-related
protease P69B. J Biol Chem 279(25):26370–26377
Tian M, Win J, Song J, van der Hooren R, van der Knaap E, Kamoun S (2007) A Phytophthora
infestans cystatin-like protein targets a novel tomato papain-like apoplastic protease. Plant
Physiol 143:364–377
Tucker SL, Talbot NJ (2001) Surface attachment and pre-penetration stage development by plant
pathogenic fungi. Annu Rev Phytopathol 39:385–417
van den Burg HA, Harrison SJ, Joosten MH, Vervoort J, de Wit PJ (2006) Cladosporium fulvum
Avr4 protects fungal cell walls against hydrolysis by plant chitinases accumulating during
infection. Mol Plant Microbe Interact 19:1420–1430
van der Hoorn RAL, Kamoun S (2008) From guard to decoy: a new model for perception of plant
pathogen effectors. Plant Cell 20:2009–2017
van der Linde K, Kastner C, Kumlehn J, Kahmann R, Doehlemann G (2010) Systemic virus
induced gene silencing allows functional characterization of maize genes during the biotrophic
interaction with Ustilago maydis. New Phytologist 189(2):471–483
van Esse HP, Bolton MD, Stergiopoulos I, de Wit PJ, Thomma BP (2007) The chitin-binding
Cladosporium fulvum effector protein Avr4 is a virulence factor. Mol Plant Microbe Interact
20:1092–1101
van Esse HP, Van’t Klooster JW, Bolton MD, Yadeta KA, van Baarlen P, Boeren S, Vervoort J, de
Wit PJ, Thomma BP (2008) The Cladosporium fulvum virulence protein Avr2 inhibits host
proteases required for basal defense. Plant Cell 20:1948–1963
238
K. Schipper and G. Doehlemann
van Kan JA (2006) Licensed to kill: the lifestyle of a necrotrophic plant pathogen. Trends Plant Sci
11:247–253
Wahl R, Wippel K, Goos S, K€amper J, Sauer N (2010a) A novel high-affinity sucrose transporter is
required for virulence of the plant pathogen Ustilago maydis. PLoS Biol 8:e1000303
Wahl R, Zahiri A, K€amper J (2010b) The Ustilago maydis b mating type locus controls hyphal
proliferation and expression of secreted virulence factors in planta. Mol Microbiol 75(1):208–220
Waller JM, Bigger M, Hillocks RJ (2007) Coffee pests, diseases and their management. CABI,
Wallingford, Oxfordshire, p 171
Walton JD (1996) Host-selective toxins: agents of compatibility. Plant Cell 8:1723–1733
Wasternack C (2007) Jasmonates: an update on biosynthesis, signal transduction and action in
plant stress response, growth and development. Ann Bot 100:681–697
Wenzler H, Meins F (1987) Persistent changes in the proliferative capacity of maize leaf tissues
induced by Ustilago infection. Physiol Mol Plant Pathol 30:309–319
Westerink N, Roth R, van den Burg HA, De Wit PJ, Joosten MH (2002) The AVR4 elicitor protein
of Cladosporium fulvum binds to fungal components with high affinity. Mol Plant Microbe
Interact 15:1219–1227
Whisson SC, Boevink PC, Moleleki L, Avrova AO, Morales JG, Gilroy EM, Armstrong MR,
Grouffaud S, van West P, Chapman S, Hein I, Toth IK, Pritchard L, Birch PR (2007) A
translocation signal for delivery of oomycete effector proteins into host plant cells. Nature
450:115–118
W€
osten HA, Bohlmann R, Eckerskorn C, Lottspeich F, B€
olker M, Kahmann R (1996) A novel
class of small amphipathic peptides affect aerial hyphal growth and surface hydrophobicity in
Ustilago maydis. EMBO J 15:4274–4281
Zahiri A, Heimel K, Wahl R, Rath M, K€amper J (2010) The Ustilago maydis forkhead transcription factor Fox1 is involved in the regulation of genes required for the attenuation of plant
defenses during pathogenic development. Mol Plant Microbe Interact 23:1118–1129
Zarnack K, Maurer S, Kaffarnik F, Ladendorf O, Brachmann A, K€amper J, Feldbr€
ugge M (2006)
Tetracycline-regulated gene expression in the pathogen Ustilago maydis. Fungal Genet Biol
43:727–738
Zheng Y, Kief J, Auffarth K, Farfsing JW, Mahlert M, Nieto F, Basse CW (2008) The Ustilago
maydis Cys2His2-type zinc finger transcription factor Mzr1 regulates fungal gene expression
during the biotrophic growth stage. Mol Microbiol 68:1450–1470
Zipfel C, Robatzek S, Navarro L, Oakeley EJ, Jones JD, Felix G, Boller T (2004) Bacterial disease
resistance in Arabidopsis through flagellin perception. Nature 428:764–767
Zipfel C, Kunze G, Chinchilla D, Caniard A, Jones JD, Boller T, Felix G (2006) Perception of the
bacterial PAMP EF-Tu by the receptor EFR restricts Agrobacterium-mediated transformation.
Cell 125:749–760
Compatible Plant-Root Knot Nematode
Interaction and Parallels with Symbiosis
Bruno Favery, Michaël Quentin, and Pierre Abad
Abstract Among plant pathogens, the root-knot nematodes (RKN), Meloidogyne
spp., are obligate biotrophic pathogens that can establish and maintain an intimate
relationship with their host plants. They are able to induce the redifferentiation of
root cells into hypertrophied and multinucleate feeding cells essential for their
development. Hyperplasia of the surrounding feeding cells lead to the organogenesis of a typical root gall. In this chapter, we describe the complex interactions
between RKN and their infected hosts. We highlight the progress in our understanding of host plant response during the compatible interaction focusing on key
plant functions involved in giant cell ontogenesis. Throughout, parallels with
symbiotic rhizobia–legume interactions are emphasized.
1 Introduction
Plants associate with a wide range of mutualistic and parasitic organisms, ranging
from bacteria to nematodes. Obligate biotrophic plant pathogens develop stable,
compatible associations with their hosts. A common feature in biotrophy is the
development of specialized interfaces between the microorganisms and the plant
cell over which nutrients are transferred (Harrison 1999). Therefore, it is quite
conceivable that these interactions might have evolved certain common core
components affecting cellular functions such as cell wall reorganization, membrane
B. Favery • M. Quentin • P. Abad (*)
Interactions Biotiques et Santé Végétale, INRA, UMR 1301, 400 route des Chappes,
06903 Sophia Antipolis, France
Interactions Biotiques et Santé Végétale, CNRS, UMR 6243, 400 route des Chappes,
06903 Sophia Antipolis, France
Interactions Biotiques et Santé Végétale, INRA-CNRS-Université de Nice Sophia-Antipolis,
UMR 1301-6243, 400 route des Chappes, F-06903 Sophia Antipolis, France
e-mail: [email protected]; [email protected]; [email protected]
S. Perotto and F. Baluska (eds.), Signaling and Communication in Plant Symbiosis,
Signaling and Communication in Plants 11, DOI 10.1007/978-3-642-20966-6_10,
# Springer-Verlag Berlin Heidelberg 2012
239
240
B. Favery et al.
synthesis, metabolite fluxes or cytoskeleton rearrangements (Parniske 2000; Lipka
and Panstruga 2005). It is a fact that the development and physiology of biotrophic
interactions are specific and significantly different from each other (depending on
the type of microorganism), but the ones occurring between plants and sedentary
endoparasitic nematodes such as RKN, and the legume–Rhizobium symbioses are
both leading to the formation of new root structures: the nodules on legume roots
induced by symbiotic nitrogen-fixing bacteria and galls containing nematode feeding cells formed by RKN (Fig 1a). As in lateral root formation, both structures are
initiated in the differentiated root zone and involve a reactivation of the cell cycle
and a subsequent redifferentiation process (Grunewald et al. 2009b). Therefore,
questions have been raised about molecular mechanisms and developmental
pathways in the plant that ultimately determine the different outcomes of these
interactions. Do these mechanisms share some common features or are they distinct
phenomena? What are the processes that determine mutualism versus parasitism?
This chapter reviews recent molecular insights into susceptible plant responses to
RKN infection and comparisons between these two processes in legumes.
Fig. 1 Cellular responses to RKN infection. (a) Galls on tomato roots infected by M. incognita.
(b) Infection of Arabidopsis root expressing MAP65-3:GFP by M. incognita J2 larvae. (c)
Selection of root vascular cells and puncture by the J2 stylet. This cell will undergo a first nuclear
division without cytokinesis leading to the formation of a binucleate cell. (d) Multinucleate and
hypertrophied giant cells. Mini cell plates (black arrow) are visible. Sections through a gall at
15 days post-infection stained with toluidine blue. Asterisks, giant cell; N nematode; Bar ¼ 10 mm
(c) or 100 mm (d)
Compatible Plant-Root Knot Nematode Interaction and Parallels with Symbiosis
241
2 RKN Infection and Giant Cell Ontogenesis
In contrast to rhizobia which interact with legume species, the RKN host range
encompasses nearly all flowering plants, leading to extensive crop loss (Abad and
Williamson 2010). The wide host range (thousand of plant species) is associated
with a worldwide distribution in all temperate and tropical areas making them
extremely successful and damaging parasites (Trudgill and Blok 2001). RKN infect
roots as microscopic vermiform second-stage juveniles (J2). Mobile J2s penetrate
the root usually at the root elongation zone (Fig. 1b) and migrate between cells to
reach the root apex and then enter into the plant vascular cylinder. Once in the stele,
RKN select five to seven root parenchyma cells (Fig. 1c) and induce their redifferentiation into elaborate feeding cells, from which RKN puncture cytoplasmic
contents with their protractible stylet (Fig. 1c). These multinucleate and
hypertrophied feeding cells, named giant cells, result from synchronous nuclear
divisions in the absence of cell division (Jones and Payne 1978). Recently, in vivo
confocal microscopy revealed that failure to form functional cell plates following
nuclear divisions may be attributed to a restricted outgrowth of the phragmoplast
microtubule array that leads to the formation of particular mini cell plates (Fig. 1d)
that do not extend across adjacent faces of the giant cell (Caillaud et al. 2008c).
Giant cells continue to enlarge for 2–3 weeks, occupying the most part of the central
cylinder. Mature giant cells may reach 100 times the size of a normal root vascular
parenchyma cells and can contain up to 100 nuclei. Nuclei are highly amoeboid and
show dispersed chromatin reflecting intensive gene transcription, as expected for an
actively growing cell. Cytoplasmic content of giant cells increase greatly and
normal vacuolization is lost. Wall ingrowths develop in contact with the xylem
elements and an extensive wall labyrinth forms, increasing the exchange surface
area at the associated membrane. These feeding cells function as specialized sinks
supplying the nematode with nutrients throughout its life cycle. After 3–8 weeks,
the pear-shaped female produces eggs that are released on the root surface. At the
onset of giant cell induction, hyperplasia is observed where surrounding parenchyma vascular cells and pericycle cells divide to form the typical galls (Fig. 1a),
which give RKN its name.
It is not yet understood how giant cells are induced, but it is certainly triggered
by nematode secretions (Davis et al. 2004; Vanholme et al. 2004). Because RKN
have such a large host range, they probably are able to hijack fundamental host
functions. The recent characterization of a RKN secreted peptide able to interact
with plant SCARECROW-like transcription factors gave new insights into the way
nematodes can manipulate plant physiology to their own benefit (Huang et al.
2006). Full genome sequencing of two RKN provides new opportunities for studying plant–nematode interactions (Bird et al. 2009). Striking similarities between
RKN and bacteria proteins have led to the discovery that some nematode parasitism
proteins such as cell wall-degrading or -modifying enzymes, usually absent from
animals, have been acquired by multiple independent lateral gene transfers from
different bacterial sources (Scholl et al. 2003; Danchin et al. 2010).
242
B. Favery et al.
3 Molecular Plant Response to RKN Infection
The routes by which nematodes manipulate their plant hosts to induce feeding cells
are still not well understood. In the past few years, molecular and genetic approaches
allowed substantial progress in our understanding of the host plant response during
the compatible interaction. These approaches were based on expression analyses of
candidate genes by promoter reporter fusions, in situ hybridization or RT-PCR, on
promoter trap and on differential gene expression between healthy and infected root
regions. The development of plant microarrays has made possible to generate largescale patterns of plant gene expression during giant cell formation. Genome-wide
expression profiling has been used to study the response of Arabidopsis thaliana or
tomato to RKN infection (Bar-Orl et al. 2005; Hammes et al. 2005; Jammes et al.
2005; Fuller et al. 2007; Bhattarai et al. 2008). These studies identified a large
number of genes being regulated in response to RKN infection, reflecting the
complexity of nematode feeding site ontogenesis (Gheysen and Fenoll 2002; Bird
and Kaloshian 2003; Caillaud et al. 2008b). More recently, material from giant cells
was captured using laser microdissection at early stages following infection,
showing that by 3 days post-inoculation (dpi) giant cells exhibit major gene regulation. This study confirmed the molecular distinctiveness of the giant cells within the
gall and showed that most of the differentially regulated genes have no previously
assigned function (Barcala et al. 2010).
These molecular approaches highlighted changes of some key plant processes
such as cytoskeleton organization and cell cycle regulation (de Almeida Engler et al.
1999; de Almeida Engler et al. 2004; Favery et al. 2004; Caillaud et al. 2008c;
Clement et al. 2009), extensive cell wall modification (Goellner et al. 2001;
Vercauteren et al. 2002; Mitchum et al. 2004), hormone and defence responses
(Lohar et al. 2004; Jammes et al. 2005; Karczmarek et al. 2008) and major
reprogramming of plant metabolism (Wang et al. 2003; Hammes et al. 2006)
(Table 1). Characterization of the genes specifically regulated during giant
cell development is a first step towards understanding compatible plant–RKN
interactions. Extensive analysis must be coupled with a detailed cellular expression
pattern study, the characterization of knockout mutants and biochemical
investigations, to more accurately dissect gene function during giant cell development. Up to date few mutants showing impaired nematode infection were
characterized (Table 1). Recently, unique defects in giant cell ontogenesis were
described in absence of regulators of microtubule (MT) or microfilament (MF)
dynamics highlighting the importance of changes in the cytoskeleton architecture
for a proper giant cell development (Caillaud et al. 2008c; Clement et al. 2009).
4 Cytoskeleton Organization
The plant cytoskeleton is a tremendously dynamic and flexible intracellular scaffold
composed mainly of MTs and actin filaments (F-actin or MFs). It plays a central
role in intracellular transports, cell division, cell differentiation and morphogenesis,
PRL
At
ADF3
ADF4
ACT2
ACT7
MAP65-3
TUB1
Tubulin b, a
At
At
At
At
At
At
At
Mt
CCS52a
Cytoskeleton
At
AtFH6
At
AtFH10
At
ADF2
Cell cycle regulator
Cell cycle regulator
Cell cycle regulator
CDC2b
CYCA2;1
CYCB1;1
At
At
At
Formin
Formin
Actin depolymerizing
factor
Actin depolymerizing
factor
Actin depolymerizing
factor
Actin
Actin
Microtubule Associated
Protein
b-tubulin
Tubul ns
DNA replication
initiation
Endoreduplication
Cell cycle regulator
Cell cycle
At
CDKA1/CDC2a
UP
UP
UP
UP
UP
UP
UP
UP
UP
UP
UP
UP
UP
UP
UP
UP
In gallb
–
YES
YES
YES
YES
NO
–
YES
YES
YES
YES
YES
YES
YES
YES
YES
In
giant
cellsc
–
–
–
–
KO: aborted giant cells
–
KO: no effect
KO: no effect
inducible RNAi: aborted giant
cells
–
–
–
RNAi pWRK23: less galls and egg
masses
–
–
–
Table 1 Functional analysis of genes differentially expressed during RKN infection
Function
Expression
Phenotyped
Planta Gene name
Lilley et al. (2004)
de Almeida Engler et al. (2004)
de Almeida Engler et al. (2004)
de Almeida Engler et al. (2004)
Caillaud et al. (2008c)
Clement et al. (2009)
Fuller et al. (2007)
Favery et al. (2004)
Jammes et al. (2005)
Clement et al. (2009)
(continued)
Favery et al. (2002) and Koltai et al. (2001)
Niebel et al. (1996), de Almeida Engler et al.
(1999) and Van de Cappelle et al. (2008)
de Almeida Engler et al. (1999)
de Almeida Engler et al. (1999)
Niebel et al. (1996) and de Almeida Engler
et al. (1999)
Huang et al. (2003)
References
Compatible Plant-Root Knot Nematode Interaction and Parallels with Symbiosis
243
Auxin responsive
Gm
RHA1/ATRAB5A
GRP7
At
At
At
AUX1
Metabolism
Gh
LEA14
UP
Late embryogenesis
abundant protein
Small GTP binding
protein
Oleosin
UP
UP
UP
Auxin transport
UP
–
UP
–
–
–
Auxin response
Cytokinin response
Cytokinin oxidase
Ethylene response factor
Ethylene resistant
GH3/DR5
UP
UP
UP
UP
UP
UP
early
In gallb
–
NO
NO
–
YES
–
–
–
NO
YES
YES
YES
YES
YES
YES
NO
In
giant
cellsc
Expression
Cellulase
Pectin acetylesterase
Cell wall protein
1-4 endoglucanase
1-4 endoglucanase
1-4 endoglucanase
Expansin
Function
Cell wall
At
CEL1
At
PAE
Np
Extensin
Nt
NtCEL7, 8
Nt
NtCEL7
Nt
NtCEL2
Sl
EXPA5
Hormone response
Sl
DIAGEOTROPICA
At
ARR5
At/Zm AtCKX3/ZmCKX1
Mt
ERF1.1
Lj
ETR1
Table 1 (continued)
Planta Gene name
–
–
–
–
KO: more resistant
–
OE: 50% less galls
OE: no effect
Transgenics carrying the
Arabidopsis dominant etr1.1:
No effect
–
–
–
–
–
–
AS lines: less egg masses
Phenotyped
Karimi et al. (2002)
Vercauteren et al. (1998)
Gheysen et al. (1996)
Hutangura et al. (1999) and Karczmarek et al.
(2004)
Mazarei et al. (2003)
Richardson and Price (1984)
Lohar et al. (2004)
Lohar et al. (2004)
Anderson et al. (2010)
Lohar and Bird (2003)
Mitchum et al. (2004)
Vercauteren et al. (2002)
Niebel et al. (1993)
Goellner et al. (2001)
Wang et al. (2007)
Goellner et al. (2001)
Gal et al. (2006)
References
244
B. Favery et al.
ATAO1
HEMOGLOBIN
mUCP
Diamine oxidase
Oxygen transport
Mitochondrial
uncoupling protein
Sl
LEMMI9
Late embryogenesis
abundant protein
Ha
HSP17.7G4
Heat shock protein
Ha
HSP17.6G2
Heat shock protein
At
RPL16A
Ribosomal protein
At
PAG1
20S proteasome subunit
At
RPE
Ribulose phosphate
epimerase
At
PGM
Phosphoglycerate
mutase
Sl
HMG2
Hydroxy-methylglutaryl
CoA reductase
At
Endomembrane protein
At
DapDC
Diaminopimelate
decarboxylase
Wound response and defense
At
PAL1
Phenylalanine
ammonia-lyase
St
GST1
Glutathione-Stransferase
St
WUN1
Wound response
Sl
TSW12
Lipid Transfer Protein
At
POX
Peroxidase
At
TPI
Trypsin protease
inhibitor
At
Pa
At
–
–
–
–
YES
YES
–
YES
YES
–
YES
YES
YES
–
YES
YES
YES
UP
UP
UP
UP
DOWN NO
–
UP
UP
UP
UP
UP
UP
UP
UP
UP
UP
–
–
–
–
–
–
–
–
–
–
KO: no galls
–
YES
UP
–
–
–
–
–
YES
UP
UP
UP
Hansen et al. (1996)
Gheysen and Fenoll (2002)
Vercauteren et al. (2001)
Vercauteren et al. (2001)
Strittmatter et al. (1996)
Goddijn et al. (1993)
Vercauteren et al. (2001)
Vercauteren et al. (2001)
Cramer et al. (1993)
Mazarei et al. (2003)
(continued)
Escobar et al. (2003) and Barcala et al. (2008)
Barcala et al. (2008)
Lilley et al. (2004)
Vercauteren et al. (2001)
Favery et al. (1998)
Escobar et al. (1999)
Møller et al. (1998)
Ehsanpour and Jones (1996)
Vercauteren et al. (2001)
Compatible Plant-Root Knot Nematode Interaction and Parallels with Symbiosis
245
TPI
COMT1
OMT
LTP
PR1
PR2
PR3
PR4
PR5
CHS1–HS3
DFG (tt3)
CHS (tt4)
CHS
CHI (tt5)
F30 H (tt7)
pC4H-F5H
LOX
PAP2
At
At
Nt
At
At
At
At
At
At
Tr
At
At
Mt
At
At
At
Zm
Nt
Table 1 (continued)
Planta Gene name
Chalcone isomerase
Flavonoid 30
hydroxylase
Ferulic acid 5hydroxylase
Lipoxygenase
MYB transcription
factor
Trypsin protease
inhibitor
Caffeate Omethyltransferase
O-methyl transferase
Lipid Transfer Protein
Pathogenesis-related
protein
b-1,3-glucanase
Endo-chitinase
Endo-chitinase
Thaumatin-like
Chalcone synthase
Dihydroflavono
reductase
Chalcone synthase
Chalcone synthase
Function
–
–
–
–
–
–
–
–
–
–
–
UP
–
–
UP
–
–
–
–
–
–
–
–
–
–
–
–
YES
–
–
–
–
YES
Wuyts et al. (2006)
Wuyts et al. (2006)
Wuyts et al. (2006)
Wuyts et al. (2006)
Wasson et al. (2009)
Hamamouch et al. (2010)
Hamamouch et al. (2010)
Hamamouch et al. (2010)
Hamamouch et al. (2010)
Hutangura et al. (1999)
Wuyts et al. (2006)
Wuyts et al. (2006)
Fuller et al. (2007)
Hamamouch et al. (2010)
Quentin et al. (2009)
Jammes et al. (2005)
References
KO: egg number increase
Gao et al. (2008)
OE: gall and egg number increase Wuyts et al. (2006)
OE: less females
KO: no effect
RNAi: reduced gall size and cell
number per gall
KO: no effect
KO: no effect
OE: no effect
OE: no effect
OE: no effect
OE: no effect
–
KO: no effect
AS lines: less egg masses
–
OE: less galls
KO: no effect
KO: no effect
In
giant
In gallb cellsc
DOWN NO
UP
Phenotyped
Expression
246
B. Favery et al.
PAL
Amino acid transporter
Amino acid transporter
Ammonium transporter
Tonoplast aquaporin
Plasma membrane
aquaporin
Serine/threonine kinase
Tonoplast aquaporin
Phenylalanine
ammonia-lyase
UP
UP
UP
UP
DOWN
DOWN
UP
–
–
YES
YES
YES
NO
NO
YES
–
–
–
–
KO: no effect
–
–
–
OE: no effect
Lilley et al. (2004)
Opperman et al. (1994)
Hammes et al. (2005)
Hammes et al. (2006)
Fuller et al. (2007)
Goddijn et al. (1993)
Hammes et al. (2005)
Wuyts et al. (2006)
Repetitive proline-rich UP
NO
OE: gall number increase
Favery et al. (2002)
proteins early
nodulin
Mt
ENOD11
Early nodulin
UP
NO
–
Boisson-Dernier et al. (2005)
At
LBD41
Lateral organ boundary UP
–
–
Fuller et al. (2007)
domain
Mt
KNOX
Transcription factor
UP
YES
–
Koltai et al. (2001)
Mt
PHAN
Transcription factor
UP
YES
–
Koltai et al. (2001)
Lj
HAR1
Clavata-like LRR–
–
KO: gall number increase
Lohar and Bird (2003)
RLKinase
At
SRS5
Short internode (SHI)- UP
YES
–
Barcala et al. (2010)
related
At
WRKY23
WRKY transcription
UP
YES
KO: no effect
Barthels et al. (1997) and Grunewald et al.
(Att0001)
factor
(2008)
Are indicated genes for which data on expression were obtained/confirmed at the cellular level using GUS- or GFP-reporter gene or RNA in situ hybridization
or for which knock-out, -down or overexpression were studied. aAt Arabidopsis thaliana; Gh Gossypium hirsutum; Gm Glycine max; Lj Lotus japonicus; Mt
Medicago truncatula; Np Nicotiana plumbaginifolia; Nt Nicotiana tabacum; Pa Parasponia andersonii; St Solanum tuberosum; Sl Solanum lycopersicum; Tr
Trifolium repens; Zm Zea mays. bUP means upregulated; DOWN downregulated gene expression. cGrey shade indicate data on the subcellular localization of
the protein in giant cells. dKnock-out (KO), anti-sense expression (AS) or overexpression (OE) effects are indicated; –, not determined
At
ARSK1
Nt
TobRB7
Miscellaneous
Mt
ENOD40
Transport
At
AAP6
At
CAT6
At
AMT1;2
At
a-TIP1-1
At
PIP2.5
Nt
Compatible Plant-Root Knot Nematode Interaction and Parallels with Symbiosis
247
248
B. Favery et al.
and is essential in regulating establishment of plant–pathogen interactions and
symbiosis. Immunostaining and use of fluorescent markers for observation of the
cytoskeleton structures within the giant cells revealed major rearrangements of both
MFs and MTs during nematode feeding sites formation (de Almeida Engler et al.
2004; Caillaud et al. 2008a; de Almeida Engler et al. 2010). Chemical blocking of
the actin or MT cytoskeleton dynamics (stabilization or breakdown) during feeding
cell initiation resulted in the arrest of proper giant cell development and consequently nematode development (Wiggers et al. 2002; de Almeida Engler et al.
2004). Microtubule-associated proteins (MAPs) and acting-binding proteins
(ABPs) play essential roles in controlling cytoskeleton dynamics and organization.
Proteins implicated in giant cell actin and MT cytoskeleton reorganization were
identified. Formins and actin-depolymerising factors (ADFs) are major regulator of
actin dynamics and architecture. Three genes coding for Arabidopsis formins
AtFH6, AtFH1 and AtFH10 were upregulated in galls upon M. incognita infection,
and AtFH6 was shown to be anchored and uniformly distributed throughout the
giant cell plasma membrane (Favery et al. 2004). It is hypothesized that formins
regulate assembly of actin cables guiding the vesicle trafficking needed for extensive plasma membrane and cell wall biogenesis and therefore giant cells isotropic
growth. Recently, ADF2 has been shown to be instrumental to the successful RKN
infection. Knocking down AtADF2 resulted in a decrease in F-actin turnover
responsible for the stabilization and bundling of the MFs. This actin cytoskeleton
stabilization did not allow nematodes to mature into females (Clement et al. 2009).
A unique defect in giant cell formation in a map65 Arabidopsis mutant was
described (Caillaud et al. 2008c). Detailed functional analyses of MAP65-3 showed
that it plays a key role in the organization of MT arrays during both mitosis (spindle
morphogenesis) and cytokinesis (phragmoplast expansion) and was associated with
mini cell plates (Fig. 1d) formed between daughter nuclei during cytokinesis
initiation in developing giant cells. In the absence of MAP65-3, giant cells started
to develop but accumulation of mitosis defects (cell wall stubs and connected
nuclei) during repeated nuclear division prevented the development of functional
feeding cells. Giant cells did not complete their differentiation process and were
eventually destroyed impairing the maturation of the infecting nematodes.
While cytoskeleton reassembly during symbiotic interactions has been studied
extensively (Takemoto and Hardham 2004; Schmidt and Panstruga 2007), identification of components regulating this process is at its beginning. The two proteins
NAP1 and PIR1 of the SCAR/WAVE complex responsible for the activation of the
ARP2/3 complex involved in actin nucleation and polymerization are the only
regulators that have been identified as being directly involved in the invasion of
Lotus japonicus roots by Mesorhizobium loti (Yokota et al. 2009).
5 Cell Cycle Regulation and Endoreduplication
The first sign characteristic of giant cell induction is the cell cycle (re)activation in
parenchyma vascular cells. Giant cells will then be subjected to synchronous
repeated mitosis without complete cytokinesis. Nodule formation is initiated by
Compatible Plant-Root Knot Nematode Interaction and Parallels with Symbiosis
249
dedifferentiation of root cortical cells followed by cell proliferation, establishing a
cluster of meristematic cells that give rise to the nodule primordium (Patriarca et al.
2004; Crespi and Frugier 2008). To initiate organogenesis, both nematodes and
symbiotic bacteria recruit cell cycle regulators, such as cyclins or cyclin-dependent
kinases, at early stages of the interactions (Table 1) (de Almeida Engler et al. 1999;
Barcala et al. 2010; Maunoury et al. 2010). In nodule, following division, cells
undergo endoreduplication – repeated cycles of DNA replication without mitosis –
allowing their differentiation into enlarged symbiotic cells (Vinardell et al. 2003).
Recently, Maunoury et al. (2010) hypothesized that it is endoreduplication that may
act as a transcriptome switch for the cell to evolve from progenitor primordium or
meristematic cell to a symbiotic cell. High expression of CCS52 was observed in
endoreduplicating nodule tissues of M. truncatula (Cebolla et al. 1999). CCS52
promotes endoreduplication by activation of the anaphase-promoting complex
(APC) resulting in mitotic cyclin destruction and mitosis arrest. Upregulation of
CCS52 was also observed in giant cells and their surrounding cells (Koltai et al.
2001; Favery et al. 2002) suggesting that endoreduplication process occurs in giant
cells. Its function still remains unclear but should represent an important process in
the development of functional giant cells to support the enhanced metabolic
demands imposed by RKN. More generally host endoreduplication is an emerging
theme in biotroph–plant interactions (Wildermuth 2010).
6 Plant Hormone Response
Since plant organogenesis is largely regulated by plant hormones, researches
have focused on whether microorganisms directly or indirectly manipulate the
plant hormone balance to induce root nodule or gall formation. Using an auxinresponsive promoter GH3 (or its synthetic derivative DR5) fused to the reporter
gene GUS, an early and transient increase in auxin response was detected in giant
cells and in white clover nodule primordia (Mathesius et al. 1998; Hutangura et al.
1999). Nodule initiation in indeterminate legumes was preceded by an inhibition of
acropetal auxin transport. Similarities in the proteomes of root cells during the early
step of nodulation and after auxin application suggest that auxin mediates substantial protein changes in nodulation initiation (van Noorden et al. 2007). Interestingly,
the M. truncatula supernumerary nodules mutant sunn presents an altered auxin
level that correlates with nodules number and protein accumulation. During RKN
infection, auxin accumulation was suggested to trigger gall formation and was
associated with an induction of the flavonoid pathway as observed in rhizobia
nodules. Several flavonoids are known to be putative auxin transport inhibitors.
However, if flavonoid-deficient roots did not form nodules they showed no difference in the number of galls. Galls induced on flavonoid-deficient roots formed
normal giant cells, but were shorter and characterized by reduced numbers of
dividing surrounding cells (Wasson et al. 2009).
250
B. Favery et al.
The increased local auxin response in neoformed organs can result from
increased auxin influx and/or an inhibition of auxin efflux. The induction observed
in microarray studies (Hammes et al. 2005; Jammes et al. 2005) of two Arabidopsis
genes coding for the auxin influx carriers AUX1 and LAX3 is consistent with a role
for auxin in the establishment of RKN. During nodule initiation a functional auxin
transport system is required since silencing of several PIN genes result in a reduced
nodulation in M. truncatula (Huo et al. 2006). How auxin importers and exporters
are regulated during nodulation and gall formation is still unknown (Grunewald
et al. 2009b). The first study on the role of the auxin transport during plant–parasitic
nematode interaction was done on cyst nematodes, another group of sedentary
parasitic nematode. Manipulation of both the expression and the subcellular polar
localization of PIN auxin transporters by cyst nematodes facilitates the initiation
and development of their multinucleate feeding sites, named syncytia (Grunewald
et al. 2009a). RKN giant cells differ significantly from syncytia that result from cell
wall dissolution and integration of surrounding cells. Moreover, in contrast to cyst
nematodes which induce significantly less infection on several auxin signalling
mutants (Goverse et al. 2000; Grunewald et al. 2009a), only one hormone mutant,
the tomato diageotropica, has been reported to alter RKN parasitism (Richardson
and Price 1984). As auxin is an important player in plant organogenesis it is likely
that auxin and auxin transport are targeted by RKN.
Not only changes in the host auxin would be required for nodules and giant cells/
gall induction but also cytokinins have emerged as an important player in these two
interactions. Using the Arabidopsis cytokinin-responsive ARR5 gene as a marker, a
role for cytokinin has been highlighted during the first steps of nodule initiation and
gall formation. This gene is, however, repressed in giant cells and in mature gall and
determinate nodules. Thus, cytokinins can play a role either as stimulator or
inhibitor of cell division. In addition, decreasing cytokinin levels by expressing a
cytokinin oxidase (CKX) resulted in a reduction of both nodules and gall numbers
on transgenic hairy roots (Lohar et al. 2004). A strict correlation between an
elevated cytokinin level and KNOX gene expression has been observed, suggesting
that cytokinins may either regulate KNOX expression or be regulated by KNOX
(Bird and Kaloshian 2003). KNOX and the Myb transcription regulator PHAN are
required for normal meristem maintenance and induced in both M. truncatula galls
and nodules again illustrating similarities between these two organs.
Phytohormones contribute also to modulate defence process. The role of the
signalling molecules jasmonic acid (JA) and salicylic acid and ethylene is well
described in shoots, however much less is known about their role in roots. Recent
advances on symbiosis and nematode interactions were recently reviewed (Gutjahr
and Paszkowski 2009). Thus, tomato susceptibility to RKN would require an intact
JA signalling pathway (Bhattarai et al. 2008).
7 Defence-Related Signalling Compounds
During invasion RKN cause little damage to the root as they migrate between the
cells. Still, a rapid and transient production of reactive oxygen species (ROS), such
as superoxide anion (O2) and hydrogen peroxide (H2O2), is detected at the time of
Compatible Plant-Root Knot Nematode Interaction and Parallels with Symbiosis
251
nematode invasion. No significant oxidative burst was detectable at the time of
giant cell induction and at later time points (Melillo et al. 2006; Das et al. 2008).
ROS are also produced in host plants during interaction with rhizobia notably in the
infection process (Santos et al. 2001). So the bacteria have to protect themselves
from these defence molecules and bacterial strains with impaired capacity to
detoxify H2O2 have deficiencies in their symbiotic capacities. Despite negative
effects, ROS has been reported as signalling molecules in the control of nodulation
process, and a threshold level of ROS would be required for a harmonious nodule
development (Pauly et al. 2006). Additional major molecules involved in the
regulation of the cellular redox state, nitrogen monoxide (NO) and glutathione
(GSH) are detected in response to rhizobia and required in proper nodule development (Frendo et al. 2005; del Giudice et al. 2011). Unfortunately, the role of GSH
and NO in giant cell formation has not yet been investigated.
Microarray analysis showed that the successful establishment of RKN is
associated with the suppression of plant defence responses in A. thaliana (Jammes
et al. 2005). Most of the defence-related genes such as WRKY transcripts and genes
associated to the JA/ethylene-dependent pathways (EIN3, ERF1, PR4) were
repressed particularly at later stage of gall formation. Expression of maize
lipoxygenase LOX3 and Arabidopsis pathogenesis-related genes PR-1, PR-2, PR3
and PR5 peaked at 7 or 9 dpi, respectively, in RKN infected roots (Gao et al. 2008;
Hamamouch et al. 2010). The observed downregulation of defence genes from host
plants suggests an active modulation of the plant response by RKN as reported in
different plant–pathogen interactions (Abramovitch and Martin 2004). Defence
suppression also appears to play an important role in symbiotic plant–microbe
interactions. The NopL effector of Rhizobium sp. NGR234 suppresses PR gene
expression when expressed in tobacco or L. japonicus (Bartsev et al. 2004).
8 Additional Nodulins Involved in RKN Parasitism
Strikingly, certain early nodulin genes (ENOD) have been shown to respond to
RKN infections. ENOD40 is also stimulated in galls from M. truncatula induced by
RKN but not in giant cells (Favery et al. 2002). ENOD40 is involved in both the
initiation and the stimulation of cortical cell division for nodule formation. It may
play a similar role in stimulating the proliferation of cells around giant cells for gall
formation. In the same way, ENOD11, a cell wall proline-rich nodulin gene early
expressed during nodulation is stimulated during RKN infection (Boisson-Dernier
et al. 2005). ENOD11 expression was detected using GUS activity exclusively in
cells surrounding the developing giant cells, as it was observed for ENOD40. The
expression patterns of these two nodulins suggest a role in cell-to-cell communication events between giant cells and vascular surrounding tissues. A larger scale
comparison using macroarrays revealed that out of 192 nodule-expressed genes
only two genes, nodulin 26 and cyclin D3, were found to be upregulated upon RKN
infection (Favery et al. 2002).
252
B. Favery et al.
Intriguingly, the host perception machinery involved in bacterial nodulation
process would be also involved in RKN infection. By comparing responses of
mutants of Nod factor receptors nfr1, nfr5, and symRK and wild-type plants in
L. japonicus, the ability of RKN to establish feeding sites and reproduce has been
shown to be markedly reduced in the mutant lines (Weerasinghe et al. 2005).
In addition, root-hair waviness and branching were detected in L. japonicus
RKN-infected roots suggesting possible common signal transduction pathway
targeted by infective RKN J2 and Nod factors in developing root hairs.
9 Conclusions
There are evidences for partial overlaps in plant microbe signalling at the cellular
and molecular levels. However, comparison between galls and nodules should take
into account that both organs are complex and contain specialized zones or cells.
Transcriptomics approach combined with laser capture microdissection will greatly
refine and expand analyses of giant cells and their surrounding cells and nodule
specific zones. A first comparison between giant cells and the infection zone II of
M. truncatula nodules indicated an overlap of 10% and 35% of genes commonly
upregulated or downregulated, respectively (Damiani et al. in preparation). These
first data highlight the recruitment of common plant pathways for infection by
pathogen and symbiont as well as the involvement of processes specific for each
type of interactions. Development of integrative ‘omics’ approach including proteomics and metabolomics will reveal the concurrent similarities and differences in
plant signalling pathways leading to different organs and developmental outcomes
as nodules or roots galls. Furthermore, substantial progress will also be made in the
next years in characterizing the direct host targets of secreted RKN effectors and
studying whether small RNAs function as key player providing unique insights
into manipulated plant cellular processes during giant cell ontogenesis. Finally,
determining how a nematode selects particular root cells and modifies them to serve
as a feeding cell will enhance our understanding of plant cell development and will
contribute to offer alternative to nematicides and novel approaches requested to
protect plant against plant parasitic nematodes.
References
Abad P, Williamson VM (2010) Plant nematode interaction: a sophisticated dialogue. In: Kader J-C,
Delseny M (eds) Advances in botanical research. Academic, New York, pp 147–192
Abramovitch RB, Martin GB (2004) Strategies used by bacterial pathogens to suppress plant
defenses. Curr Opin Plant Biol 7:356–364
Anderson JP, Lichtenzveig J, Gleason C, Oliver RP, Singh KB (2010) The B-3 ethylene response
factor MtERF1-1 mediates resistance to a subset of root pathogens in Medicago truncatula
without adversely affecting symbiosis with rhizobia. Plant Physiol 154:861–873
Compatible Plant-Root Knot Nematode Interaction and Parallels with Symbiosis
253
Barcala M, Garcia A, Cubas P, Almoguera C, Jordano J, Fenoll C, Escobar C (2008) Distinct heatshock element arrangements that mediate the heat shock, but not the late-embryogenesis
induction of small heat-shock proteins, correlate with promoter activation in root-knot nematode feeding cells. Plant Mol Biol 66:151–164
Barcala M, Garcia A, Cabrera J, Casson S, Lindsey K, Favery B, Garcia-Casado G, Solano R,
Fenoll C, Escobar C (2010) Early transcriptomic events in microdissected Arabidopsis nematode-induced giant cells. Plant J 61:698–712
Bar-Orl C, Kapulnik Y, Koltai H (2005) A broad characterization of the transcriptional profile of
the compatible tomato response to the plant parasitic root knot nematode Meloidogyne
javanica. Eur J Plant Pathol 111:181–192
Barthels N, Van der Lee FM, Klap J, Goddijn OJM, Karimi M, Puzio P, Grundler FMW, Ohl SA,
Lindsey K, Robertson VM, Van Montagu M, Gheysen G, Sijmons PC (1997) Regulatory
sequences of Arabidopsis drive reporter gene expression in nematode feeding structures. Plant
Cell 9:2119–2134
Bartsev AV, Deakin WJ, Boukli NM, McAlvin CB, Stacey G, Malnoe P, Broughton WJ, Staehelin
C (2004) NopL, an effector protein of Rhizobium sp. NGR234, thwarts activation of plant
defense reactions. Plant Physiol 134:871–879
Bhattarai KK, Xie QG, Mantelin S, Bishnoi U, Girke T, Navarre DA, Kaloshian I (2008) Tomato
susceptibility to root-knot nematodes requires an intact jasmonic acid signaling pathway. Mol
Plant Microbe Interact 21:1205–1214
Bird DM, Kaloshian I (2003) Are roots special? Nematodes have their say. Physiol Mol Plant
Pathol 62:115–123
Bird DM, Williamson VM, Abad P, McCarter J, Danchin EG, Castagnone-Sereno P, Opperman
CH (2009) The genomes of root-knot nematodes. Annu Rev Phytopathol 47:333–351
Boisson-Dernier A, Andriankaja A, Chabaud M, Niebel A, Journet EP, Barker DG, de CarvalhoNiebel F (2005) MtENOD11 gene activation during rhizobial infection and mycorrhizal
arbuscule development requires a common AT-rich-containing regulatory sequence. Mol
Plant Microbe Interact 18:1269–1276
Caillaud MC, Abad P, Favery B (2008a) Cytoskeleton reorganization: a key process in root-knot
nematode-induced giant cell ontogenesis. Plant Signal Behav 3:816–818
Caillaud MC, Dubreuil G, Quentin M, Perfus-Barbeoch L, Lecomte P, de Almeida Engler J,
Abad P, Rosso MN, Favery B (2008b) Root-knot nematodes manipulate plant cell functions
during a compatible interaction. J Plant Physiol 165:104–113
Caillaud MC, Lecomte P, Jammes F, Quentin M, Pagnotta S, Andrio E, de Almeida Engler J,
Marfaing N, Gounon P, Abad P, Favery B (2008c) MAP65-3 microtubule-associated protein is
essential for nematode-induced giant cell ontogenesis in Arabidopsis. Plant Cell 20:423–437
Cebolla A, Vinardell JM, Kiss E, Olah B, Roudier F, Kondorosi A, Kondorosi E (1999) The
mitotic inhibitor ccs52 is required for endoreduplication and ploidy-dependent cell enlargement in plants. EMBO J 18:4476–4484
Clement M, Ketelaar T, Rodiuc N, Banora MY, Smertenko A, Engler G, Abad P, Hussey PJ,
de Almeida Engler J (2009) Actin-depolymerizing factor2-mediated actin dynamics are essential for root-knot nematode infection of Arabidopsis. Plant Cell 21:2963–2979
Cramer CL, Weissenborn DL, Cotttingham CK, Denbow CJ, Eisenbach JD, Radin DN, Yu X
(1993) Regulation of defense-related gene expression during plant-pathogen interactions.
J Nematol 25:507–551
Crespi M, Frugier F (2008) De novo organ formation from differentiated cells: root nodule
organogenesis. Sci Signal 1:re11
Danchin EG, Rosso MN, Vieira P, de Almeida-Engler J, Coutinho PM, Henrissat B, Abad P (2010)
Multiple lateral gene transfers and duplications have promoted plant parasitism ability in
nematodes. Proc Natl Acad Sci USA 107:17651–17656
Das S, DeMason DA, Ehlers JD, Close TJ, Roberts PA (2008) Histological characterization of
root-knot nematode resistance in cowpea and its relation to reactive oxygen species modulation. J Exp Bot 59:1305–1313
254
B. Favery et al.
Davis EL, Hussey RS, Baum TJ (2004) Getting to the roots of parasitism by nematodes. Trends
Parasitol 20:134–141
de Almeida Engler J, De Vleesschauwer V, Burssens S, Celenza JL Jr, Inze D, Van Montagu M,
Engler G, Gheysen G (1999) Molecular markers and cell cycle inhibitors show the importance
of cell cycle progression in nematode-induced galls and syncytia. Plant Cell 11:793–808
de Almeida Engler J, Van Poucke K, Karimi M, De Groodt R, Gheysen G, Engler G (2004)
Dynamic cytoskeleton rearrangements in giant cells and syncytia of nematode-infected roots.
Plant J 38:12–26
de Almeida Engler J, Rodiuc N, Smertenko A, Abad P (2010) Plant actin cytoskeleton remodelling by plant parasitic nematodes. Plant Signal Behav 5:1–5
del Giudice J, Cam Y, Damiani I, Fung-Chat F, Meilhoc E, Bruand C, Brouquisse R, Puppo A,
Boscari A (2011) Nitric oxide is required for an optimal establishment of the Medicago
truncatula–Sinorhizobium meliloti symbiosis. New Phytol 191:405–417
Ehsanpour AA, Jones MGK (1996) Glucuronidase expression in transgenic tobacco roots with a
Parasponia promoter on infection with Meloidogyne javanica. J Nematol 28:407–413
Escobar C, De Meutter J, Aristizabal FA, Sanz-Alferez S, del Campo FF, Barthels N, Van der
Eycken W, Seurinck J, van Montagu M, Gheysen G, Fenoll C (1999) Isolation of the LEMMI9
gene and promoter analysis during a compatible plant-nematode interaction. Mol Plant
Microbe Interact 12:440–449
Escobar C, Barcala M, Portillo M, Almoguera C, Jordano J, Fenoll C (2003) Induction of the
Hahsp17.7G4 promoter by root-knot nematodes: involvement of heat-shock elements in
promoter activity in giant cells. Mol Plant Microbe Interact 16:1062–1068
Favery B, Lecomte P, Gil N, Bechtold N, Bouchez D, Dalmasso A, Abad P (1998) RPE, a plant
gene involved in early developmental steps of nematode feeding cells. EMBO J 17:6799–6811
Favery B, Complainville A, Vinardell JM, Lecomte P, Vaubert D, Mergaert P, Kondorosi A,
Kondorosi E, Crespi M, Abad P (2002) The endosymbiosis-induced genes ENOD40 and
CCS52a are involved in endoparasitic-nematode interactions in Medicago truncatula. Mol
Plant Microbe Interact 15:1008–1013
Favery B, Chelysheva LA, Lebris M, Jammes F, Marmagne A, De Almeida-Engler J, Lecomte P,
Vaury C, Arkowitz RA, Abad P (2004) Arabidopsis formin AtFH6 is a plasma membraneassociated protein upregulated in giant cells induced by parasitic nematodes. Plant Cell
16:2529–2540
Frendo P, Harrison J, Norman C, Hernandez Jimenez MJ, Van de Sype G, Gilabert A, Puppo A
(2005) Glutathione and homoglutathione play a critical role in the nodulation process of
Medicago truncatula. Mol Plant Microbe Interact 18:254–259
Fuller VL, Lilley CJ, Atkinson HJ, Urwin P (2007) Differential gene expression in Arabidopsis
following infection by plant-parasitic nematodes Meloidogyne incognita and Heterodera
schachtii. Mol Plant Pathol 8:595–609
Gal TZ, Aussenberg ER, Burdman S, Kapulnik Y, Koltai H (2006) Expression of a plant expansin
is involved in the establishment of root knot nematode parasitism in tomato. Planta
224:155–162
Gao X, Starr J, Gobel C, Engelberth J, Feussner I, Tumlinson J, Kolomiets M (2008) Maize 9lipoxygenase ZmLOX3 controls development, root-specific expression of defense genes, and
resistance to root-knot nematodes. Mol Plant Microbe Interact 21:98–109
Gheysen G, Fenoll C (2002) Gene expression in nematode feeding sites. Annu Rev Phytopathol
40:191–219
Gheysen G, Van der Eychen W, Barthels N, Karimi M, Van Montagu M (1996) The exploitation of
nematode-responsive plant genes in novel nematode control methods. Pesticide Sci 47:95–101
Goddijn OJM, Lindsey K, Van der Lee FM, Klap JC, Sijmons PC (1993) Differential gene
expression in nematode-induced feeding structures of transgenic plants harbouring promotergusA fusion constructs. Plant J 4:863–873
Goellner M, Wang X, Davis EL (2001) Endo-beta-1,4-glucanase expression in compatible plantnematode interactions. Plant Cell 13:2241–2255
Compatible Plant-Root Knot Nematode Interaction and Parallels with Symbiosis
255
Goverse A, Overmars H, Engelbertink J, Schots A, Bakker J, Helder J (2000) Both induction and
morphogenesis of cyst nematode feeding cells are mediated by auxin. Mol Plant Microbe
Interact 13:1121–1129
Grunewald W, Karimi M, Wieczorek K, Van de Cappelle E, Wischnitzki E, Grundler F, Inze D,
Beeckman T, Gheysen G (2008) A role for AtWRKY23 in feeding site establishment of plantparasitic nematodes. Plant Physiol 148:358–368
Grunewald W, Cannoot B, Friml J, Gheysen G (2009a) Parasitic nematodes modulate PINmediated auxin transport to facilitate infection. PLoS Pathog 5:e1000266
Grunewald W, van Noorden G, Van Isterdael G, Beeckman T, Gheysen G, Mathesius U (2009b)
Manipulation of auxin transport in plant roots during Rhizobium symbiosis and nematode
parasitism. Plant Cell 21:2553–2562
Gutjahr C, Paszkowski U (2009) Weights in the balance: jasmonic acid and salicylic acid signaling
in root-biotroph interactions. Mol Plant Microbe Interact 22:763–772
Hamamouch N, Li C, Seo PJ, Park C-M, Davis EL (2010) Expression of Arabidopsis pathogenesis-related genes during nematode infection. Mol Plant Pathol 12:355–364
Hammes UZ, Schachtman DP, Berg RH, Nielsen E, Koch W, McIntyre LM, Taylor CG (2005)
Nematode-induced changes of transporter gene expression in Arabidopsis roots. Mol Plant
Microbe Interact 18:1247–1257
Hammes UZ, Nielsen E, Honaas LA, Taylor CG, Schachtman DP (2006) AtCAT6, a sink-tissuelocalized transporter for essential amino acids in Arabidopsis. Plant J 48:414–426
Hansen E, Harper G, McPherson MJ, Atkinson HJ (1996) Differential expression patterns of the
wound-inducible transgene wun1-uidA in potato roots following infection with either cyst or
root knot nematodes. Physiol Mol Plant Pathol 48:161–170
Harrison MJ (1999) Biotrophic interfaces and nutrient transport in plant/fungal symbioses. J Exp
Bot 50:1013–1022
Huang X, Springer PS, Kaloshian I (2003) Expression of the Arabidopsis MCM Gene
PROLIFERA during root-knot and cyst nematode infection. Phytopathology 93:35–41
Huang G, Dong R, Allen R, Davis EL, Baum TJ, Hussey RS (2006) A root-knot nematode
secretory peptide functions as a ligand for a plant transcription factor. Mol Plant Microbe
Interact 19:463–470
Huo X, Schnabel E, Hughes K, Frugoli J (2006) RNAi Phenotypes and the localization of a
protein::GUS fusion imply a role for Medicago truncatula PIN genes in nodulation. J Plant
Growth Regul 25:156–165
Hutangura P, Mathesius U, Jones MGK, Rolfe BG (1999) Auxin induction is a trigger for root gall
formation caused by root-knot nematodes in white clover and is associated with the activation
of the flavonoid pathway. Aust J Plant Physiol 26:221–231
Jammes F, Lecomte P, de Almeida-Engler J, Bitton F, Martin-Magniette ML, Renou JP, Abad P,
Favery B (2005) Genome-wide expression profiling of the host response to root-knot nematode
infection in Arabidopsis. Plant J 44:447–458
Jones MGK, Payne HL (1978) Early stages of nematode-induced giant cell formation in roots of
Impatiens balsamina. J Nematol 10:70–84
Karczmarek A, Overmars H, Helder J, Goverse A (2004) Feeding cell development by cyst and
root-knot nematodes involves a similar early, local and transient activation of a specific auxininducible promoter element. Mol Plant Pathol 5:343–346
Karczmarek A, Fudali S, Lichocka M, Sobczak M, Kurek W, Janakowski S, Roosien J,
Golinowski W, Bakker J, Goverse A, Helder J (2008) Expression of two functionally distinct
plant endo-beta-1,4-glucanases is essential for the compatible interaction between potato cyst
nematode and its hosts. Mol Plant Microbe Interact 21:791–798
Karimi M, de Oliveira Manes C-L, Van Montagu M, Gheysen G (2002) Activation of a Pgrp-gus
fusion in Arabidopsis thaliana roots upon nematode infection. J Nematol 34:75–79
Koltai H, Dhandaydham M, Opperman C, Thomas J, Bird D (2001) Overlapping plant signal
transduction pathways induced by a parasitic nematode and a rhizobial endosymbiont. Mol
Plant Microbe Interact 14:1168–1177
256
B. Favery et al.
Lilley CJ, Urwin PE, Johnston KA, Atkinson HJ (2004) Preferential expression of a plant cystatin
at nematode feeding sites confers resistance to Meloidogyne incognita and Globodera pallida.
Plant Biotechnol J 2:3–12
Lipka V, Panstruga R (2005) Dynamic cellular responses in plant-microbe interactions. Curr Opin
Plant Biol 8:625–631
Lohar DP, Bird DM (2003) Lotus japonicus: a new model to study root-parasitic nematodes. Plant
Cell Physiol 44:1176–1184
Lohar DP, Schaff JE, Laskey JG, Kieber JJ, Bilyeu KD, Bird DM (2004) Cytokinins play opposite
roles in lateral root formation, and nematode and Rhizobial symbioses. Plant J 38:203–214
Mathesius U, Schlaman HR, Spaink HP, Of Sautter C, Rolfe BG, Djordjevic MA (1998) Auxin
transport inhibition precedes root nodule formation in white clover roots and is regulated by
flavonoids and derivatives of chitin oligosaccharides. Plant J 14:23–34
Maunoury N, Redondo-Nieto M, Bourcy M, Van de Velde W, Alunni B, Laporte P, Durand P,
Agier N, Marisa L, Vaubert D, Delacroix H, Duc G, Ratet P, Aggerbeck L, Kondorosi E,
Mergaert P (2010) Differentiation of symbiotic cells and endosymbionts in Medicago
truncatula nodulation are coupled to two transcriptome-switches. PLoS One 5:e9519
Mazarei M, Lennon KA, Puthoff DP, Rodermel SR, Baum TJ (2003) Expression of an Arabidopsis
phosphoglycerate mutase homologue is localized to apical meristems, regulated by hormones,
and induced by sedentary plant-parasitic nematodes. Plant Mol Biol 53:513–530
Melillo MT, Leonetti P, Bongiovanni M, Castagnone-Sereno P, Bleve-Zacheo T (2006) Modulation of reactive oxygen species activities and H2O2 accumulation during compatible and
incompatible tomato-root-knot nematode interactions. New Phytol 170:501–512
Mitchum MG, Sukno S, Wang X, Shani Z, Tsabary G, Shoseyov O, Davis EL (2004) The promoter
of the Arabidopsis thaliana Cel1 endo-1,4-ß glucanase gene is differentially expressed in plant
feeding cells induced by root-knot and cyst nematodes. Mol Plant Pathol 5:175–181
Møller SG, Urwin PE, Atkinson HJ, Mc-Pherson MJ (1998) Nematode-induced expression of
atao1, a gene encoding an extracellular diamine oxidase associated with developing vascular
tissue. Physiol Mol Plant Pathol 53:73–79
Niebel A, De Almeida Engler J, Tire C, Engler G, Van Montagu M, Gheysen G (1993) Induction
patterns of an extensin gene in tobacco upon nematode infection. Plant Cell 5:1697–1710
Niebel A, de Almeida Engler J, Hemerly A, Ferreira P, Inze D, Van Montagu M, Gheysen G
(1996) Induction of cdc2a and cyc1At expression in Arabidopsis thaliana during early phases
of nematode-induced feeding cell formation. Plant J 10:1037–1043
Opperman CH, Taylor CG, Conkling MA (1994) Root-knot nematode-directed expression of a
plant root-specific gene. Science 263:221–223
Parniske M (2000) Intracellular accommodation of microbes by plants: a common developmental
program for symbiosis and disease? Curr Opin Plant Biol 3:320–328
Patriarca EJ, Tate R, Ferraioli S, Iaccarino M (2004) Organogenesis of legume root nodules. Int
Rev Cytol 234:201–262
Pauly N, Pucciariello C, Mandon K, Innocenti G, Jamet A, Baudouin E, Herouart D, Frendo P,
Puppo A (2006) Reactive oxygen and nitrogen species and glutathione: key players in the
legume-Rhizobium symbiosis. J Exp Bot 57:1769–1776
Quentin M, Allasia V, Pegard A, Allais F, Ducrot PH, Favery B, Levis C, Martinet S, Masur C,
Ponchet M, Roby D, Schlaich NL, Jouanin L, Keller H (2009) Imbalanced lignin biosynthesis
promotes the sexual reproduction of homothallic oomycete pathogens. PLoS Pathog 5:
e1000264
Richardson L, Price NS (1984) Observations on the biology of Meloidogyne incognita and the
diageotropica tomato mutant. Rev Nématol 7:97–99
Santos R, Herouart D, Sigaud S, Touati D, Puppo A (2001) Oxidative burst in alfalfaSinorhizobium meliloti symbiotic interaction. Mol Plant Microbe Interact 14:86–89
Schmidt SM, Panstruga R (2007) Cytoskeleton functions in plant-microbe interactions. Physiol
Mol Plant Pathol 71:135–148
Compatible Plant-Root Knot Nematode Interaction and Parallels with Symbiosis
257
Scholl EH, Thorne JL, McCarter JP, Bird DM (2003) Horizontally transferred genes in plantparasitic nematodes: a high-throughput genomic approach. Genome Biol 4:R39
Strittmatter G, Gheysen G, Gianinazzi-Pearson V, Hahn K, Niebel A, Rohde W, Tacke E (1996)
Infections with various types of organisms stimulate transcription from a short promoter
fragment of the potato gst1 gene. Mol Plant Microbe Interact 9:68–73
Takemoto D, Hardham AR (2004) The cytoskeleton as a regulator and target of biotic interactions
in plants. Plant Physiol 136:3864–3876
Trudgill DL, Blok VC (2001) Apomictic, polyphagous root-knot nematodes: exceptionally successful and damaging biotrophic root pathogens. Annu Rev Phytopathol 39:53–77
Van de Cappelle E, Plovie E, Kyndt T, Grunewald W, Cannoot B, Gheysen G (2008) AtCDKA;1
silencing in Arabidopsis thaliana reduces reproduction of sedentary plant-parasitic nematodes.
Plant Biotechnol J 6:749–757
van Noorden GE, Kerim T, Goffard N, Wiblin R, Pellerone FI, Rolfe BG, Mathesius U (2007)
Overlap of proteome changes in Medicago truncatula in response to auxin and Sinorhizobium
meliloti. Plant Physiol 144:1115–1131
Vanholme B, De Meutter J, Tytgat T, Van Montagu M, Coomans A, Gheysen G (2004) Secretions
of plant-parasitic nematodes: a molecular update. Gene 332:13–27
Vercauteren I, Goeleven E, Barthels N, Van Montagu M, Gheysen G (1998) The rha1 gene,
encoding a small GTP binding protein, is induced in nematode infection sites. Arch Physiol
Biochem 106:158 (B130)
Vercauteren I, Van Der Schueren E, Van Montagu M, Gheysen G (2001) Arabidopsis thaliana
genes expressed in the early compatible interaction with root-knot nematodes. Mol Plant
Microbe Interact 14:288–299
Vercauteren I, de Almeida Engler J, De Groodt R, Gheysen G (2002) An Arabidopsis thaliana
pectin acetylesterase gene is upregulated in nematode feeding sites induced by root-knot and
cyst nematodes. Mol Plant Microbe Interact 15:404–407
Vinardell JM, Fedorova E, Cebolla A, Kevei Z, Horvath G, Kelemen Z, Tarayre S, Roudier F,
Mergaert P, Kondorosi A, Kondorosi E (2003) Endoreduplication mediated by the anaphasepromoting complex activator CCS52A is required for symbiotic cell differentiation in
Medicago truncatula nodules. Plant Cell 15:2093–2105
Wang Z, Potter RH, Jones MGK (2003) Differential display analysis of gene expression in the
cytoplasm of giant cells induced in tomato roots by Meloidogyne javanica. Mol Plant Pathol
4:361–371
Wang X, Replogle A, Davis E, Mitchum M (2007) The tobacco Cel7 gene promoter is auxinresponsive and locally induced in nematode feeding sites of heterologous plants. Mol Plant
Pathol 8:423–436
Wasson AP, Ramsay K, Jones MG, Mathesius U (2009) Differing requirements for flavonoids
during the formation of lateral roots, nodules and root knot nematode galls in Medicago
truncatula. New Phytol 183:167–179
Weerasinghe RR, Bird DM, Allen NS (2005) Root-knot nematodes and bacterial Nod factors elicit
common signal transduction events in Lotus japonicus. Proc Natl Acad Sci USA
102:3147–3152
Wiggers RJ, Thornton NT, Starr JL (2002) The effects of colchicine on number of giant cell nuclei
and nematode development in Pisum sativum infected by Meloidogyne incognita. Nematology
4:107–109
Wildermuth MC (2010) Modulation of host nuclear ploidy: a common plant biotroph mechanism.
Curr Opin Plant Biol 13:449–458
Wuyts N, Lognay G, Swennen R, De Waele D (2006) Nematode infection and reproduction in
transgenic and mutant Arabidopsis and tobacco with an altered phenylpropanoid metabolism.
J Exp Bot 57:2825–2835
Yokota K, Fukai E, Madsen LH, Jurkiewicz A, Rueda P, Radutoiu S, Held M, Hossain MS,
Szczyglowski K, Morieri G, Oldroyd GE, Downie JA, Nielsen MW, Rusek AM, Sato S, Tabata
S, James EK, Oyaizu H, Sandal N, Stougaard J (2009) Rearrangement of actin cytoskeleton
mediates invasion of Lotus japonicus roots by Mesorhizobium loti. Plant Cell 21:267–284
Index
A
Accommodation, 60
Actin, 242, 243, 248
Acting binding protein (ABP), 248
Actinomycetales, 75
Actinorhiza, 80
Actinorhizal plants
early signal molecules
Frankia signals, 81
mitotic activity in infected roots, 81–82
plant signals, 80–81
Frankia (see Frankia microorganism)
gene expression during infection, 84
heterologous promoters expression, 85
hormones
auxins, 86
cytokinin and actinorhizal nodulation,
86–87
Frankia, 85–86
nodule
development, 80
infection process, 78–79
signal transduction pathway
other candidates, 83–84
SymRK gene, 82–83
Adenylate cyclase, 106
Appressoria, 222
Appressorium, 187
Arabidopsis thaliana, 131, 132
Arabinogalactan protein (AGPs), 107
Arbuscular mycorrhizas (AM), 51
Arbuscules, 53, 63
Arum, 53
Asymbiotic phase, 52
Auxins, 63, 86, 131–136, 244, 249, 250
Azolla, 101–103, 105, 108–110, 112–115
B
Basidiomycete, 216
Biotrophic ascomycetes, 186
Biotrophic interaction, 225
Biotrophic interfacial complex, 188, 199
Biotrophy, 214
Branching factors, 129
C
Calcium, 143, 148–151, 163–165, 167, 168
signals, 59
signature, 59
spiking, 59
cAMP, 160, 163, 164
Carrot, 61
Casuarina, 73–81, 83, 86, 87
Casuarinaceae, 74, 76, 81, 86
Cell cycle, 240, 242, 243, 248–249
Cell death, 220
Cell penetrating peptides, 202
CgSymRK gene, 82–83
Chemotaxis, 99, 104–105
Classical phytohormones, 130
Clusters, 225, 226
Colonization, 226
Comparative genomics, 231
Compatibility, 214
Comptonia, 73, 74, 76, 78, 80
Confocal microscopy, 232
Crinklers, 196, 199
Cycad, 94, 96–99, 104, 106, 111–114
Cytokinin, 36, 37, 244, 250
Cytokinin and actinorhizal nodulation,
86–87
Cytoskeleton, 240, 242–248
S. Perotto and F. Baluska (eds.), Signaling and Communication in Plant Symbiosis,
Signaling and Communication in Plants 11, DOI 10.1007/978-3-642-20966-6,
# Springer-Verlag Berlin Heidelberg 2012
259
260
D
Dark brown and structures, 186–189
Defence response, 242, 251
E
Ectomycorrhiza (ECM)
development, 126, 129–136
ectomycorrhizal signalling evolution,
136–137
functioning structure, 124
fungi, 124
hormone signalling
auxin cell loosening and hartig net
development, 132–136
auxin, early contact phase, 131–132
volatile phytohormones as precontact
signals, 130–131
plant signals, 129
root cell walls remodelling, 128–129
root development, 127–128
secreted fungal molecules, 136
signal exchange, 135
Effectome, 231
Effector molecules, 60
Effectors, 192, 195, 196, 223
Effector-triggered immunity (ETI), 195
Endocytosis, 200
Endophyte, 143, 144, 146, 147, 150–153,
170–172
Endoplasmic reticulum, 60
Endoreduplication, 243, 248–249
ENOD 11, 247, 251
Enzymes, 222
Ethylene, 244, 250, 251
Extrahaustorial matrix, 190
Extrahaustorial membrane, 191
Extra-invasive hyphal membrane, 188, 193
F
Filament, 217
Flavonoids
biological activity, 3–5
structure and synthesis, 2–3
Frankia microorganism
general features, 75–76
hormones, 85–86
hyphae, 78, 80
molecular tools and genomics, 76–77
vs. Rhizobia, 74
signals, 81
specificity of symbiotic association, 76
stages of root infection, 78
Index
G
Gall, 240–252
Genome, 218
Giant cell, 240–244, 246–252
Glomeromycota, 52
Glomus intraradices, 65
Golgi, 60
G-protein, 157, 159–161, 163
G-protein coupled receptor, 157, 159–161
Gunnera, 94, 98–101, 104, 106, 107, 109,
111–115
H
Haustoria, 185, 190, 216
Haustorium, 187
Hemibiotrophy, 187, 215
Heterocyst, development, 103, 107, 108,
110–111
Histidine kinase, 155, 161–163
Hormogonia
development, 103, 104, 107–110, 115
repression, 108–110
role in plant infection, 103–106
Hormones, 63, 220, 242, 244, 249–250
auxins, 86
cytokinin and actinorhizal nodulation,
86–87
Frankia, 85–86
signalling, 130–136
Hornwort, 95–98, 106, 108–110, 112–114
Host defence, 169–170
Hypersensitive response (HR), 219
Hypervirulence, 229
Hyphae, 52
Hyphopodium, 53
I
Identification of effectors, 195
Infection, 215
crack entry, 42–43
peg entry, 43–44
root hair, 32, 37, 39–44
strategies, 184–190
Inflorescence, 230
Intercellular infection, 78–79
Interface, 53, 63, 224
Intracellular infection, 78–79
J
Jasmonic acid (JA), 250, 251
Index
L
Laccaria bicolor, 124, 127, 130–132,
135, 136
Lectin, 108
Lipochitooligosaccharide (LCO), 6, 18–19
Lipochito-oligosaccharides, 56
Liverwort, 95–97, 104, 106, 107, 112, 113
L. laccata, 131
Localization, 223
Lotus japonicus, 31–44, 55
LysM domains, 35–36
M
Magnaporthe oryzae, 187
Maize, 61
MAP kinase, 149, 151–159, 161
cell wall integrity, 152, 156–159, 169
pheromone response, 149, 153–157, 160
stress-activated, 149, 151–154, 161
Mating, 226
Medicago truncatula, 57
Meloidogyne ssp., 239, 240–253, 259–263
Meristem, 144, 147
Microarray, 219
Microfilaments (MF), 62, 242, 248
Microtubule-associated protein (MAP), 240,
243, 248
Microtubules (MT), 62, 241–243, 248
Mitosis, 248, 249
Moss, 95–96
Mycelium, 52
Myc factors, 17–18, 56
Mycorrhiza induced small secreted proteins
(MiSSPs), 136
N
NADPH oxidase (Nox), 146–149, 164
Nematode, 239–253, 259–263
Nematode secretions, 241
Nitrogen fixation, 94, 95, 102, 107–114
NodD, 33
Nod factor, 56
core structure, 7
decorations, 8, 34
perception, 13–14, 33–37, 41–43
receptors, 34–37, 40–44
secretion, 12
Nodule, 240, 248–252
determinate, 31, 32, 40
indeterminate, 32, 40
Nodulin, 247, 251–252
261
Nucleoporins, 57
Nutrients, 64
acquisition, 192
O
Obligate biotrophs, 185
Oomycete P. infestans, 223
Oomycetes, 189
Organ specificity, 229
Orobanche, 55
Overexpression, 227
P
PAMP, 223
PAMP-triggered immunity (PTI), 194
Parasitic plants, 55
Paris, 53
Pathogenesis related genes (PR), 246, 251
Pathogenicity, 231
Penetration, 215
Periarbuscular membrane, 53
Perifungal membrane, 53
Petunia, 61
Phosphate, 64
phosphatidylinositol–3-phosphate, 201
Photosynthesis, 221
Phytohormones, 129–132
Pili, 104
Pisolithus tinctorius, 128–131, 134
Plant colonization, 216
Plant defense, 213
Polarisation, 62
Powdery mildews, 186
PpGH3–16 gene, 133
PpIAA88 gene, 133
Prepenetration apparatus (PPA), 60
Presymbiotic phase, 53
Primary intracellular hyphae, 189
Promoter, 228
Pseudonodules, 86
PtdIns(3)P, 201
R
Reactive oxygen species (ROS), 250, 251
Receptor-like kinase, 57
Regulation, 226
Reprogramming, 230
Resistance (proteins), 224
RGD, 202
RNA sequencing (RNAseq), 169, 172
262
Root cell walls, 128–129
Root development, 127–128
Root-knot nematode (RKN), 239–252
Rust fungi, 185
Rusts, 215
RXLR, 195, 199
S
Secondary hyphae, 189
Secondary metabolite, 171, 172
Secreted proteins, 228
Secreted small proteins (SSP), 136
Secretion, 215, 222, 224
Sequencing, 225
Signalling
cross, 36, 41
cytokinin, 36, 37
Nod factor, 34, 37, 41, 43
parallel, 40–42
Smut, 216
Spores, 52
Sporidia, 217, 218
Starch, 56
Striga, 55
Index
Strigolactones, 55
biological activity, 15–17
structure and synthesis, 15
Symbiosis related acidic polypeptides
(SRAPs), 128
Symbiotic phase, 53
SYM genes, 57
SymRK gene, 82–83
T
Tassels, 230
Tomato, 61
Transcription factor, 226
Transporter, 221
Tumor, 230
U
Uredospore, 185
V
Vicia faba, 63
Virulence factors, 218